INTERACTION OF HERG CHANNELS AND SYNTAXIN 1A

by

Anton Mihic

A thesis submitted in conformity with the requirements for the degree of Master of Science Graduate Department of Physiology University of Toronto

© Copyright by Anton Mihic (2009) Interaction of hERG Channels and Syntaxin 1A

Anton Mihic

Master of Science

Graduate Department of Physiology

University of Toronto

2009

Abstract

The human ether-à-go-go related gene (hERG) encodes the pore-forming voltage-gated K+ channel that is essential for cardiac repolarization. Dr. Tsushima’s laboratory has previously characterized the endogenous expression of SNARE proteins in the mammalian heart, and the interaction of the SNARE protein syntaxin 1A (STX1A) with several cardiac ion channels. Here, we utilize a multi-disciplinary approach to describe the inhibitory effect of STX1A on hERG channel function. STX1A impairs hERG channel maturation and trafficking to the plasma membrane and induces a hyperpolarizing shift in the voltage-sensitivity of steady-state inactivation. We identify the residues involved in this protein- protein interaction through the use of hERG truncation mutations. We also describe the pharmacological and temperature-mediated rescue of hERG channel trafficking in the presence of

STX1A. The regulation of cardiac ion channels by SNARE proteins represents a novel biological mechanism that may have universally intrinsic implications for normal and diseased heart function.

-ii- Acknowledgments

I would like to thank my supervisory committee members Dr. Lyanne Schlichter and Dr. Peter Pennefather who provided valuable insight and objective feedback throughout my studies as a graduate student. Thank you for your support and encouragement. Many thanks as well to the members of my examination committee: Dr. Scott Heximer, Dr. Zhong-Ping Feng, Dr. Vijay Chauhan and Dr. Lyanne Schlichter. Additionally, I would like to acknowledge and thank our collaborators Dr. Herbert Gaisano and Dr. Alvin Shrier, whose insightful and enthusiastic input into this project has made it even more intellectually fruitful.

Completion of this thesis would not have been possible without the help of my lab mates, both past and present that tirelessly helped me to learn and refine new techniques at the lab bench. Special thanks go to the people who supported me throughout this learning process, especially Xiaodong Gao, Dr. Fuzhen Xia, Dr. Yukman Leung, Tom Zhao and Andrew Cooper.

Thank you to my parents for fostering an interest in science and discovery from an early age and for pushing me to enroll at the University of Toronto as an undergraduate student. Thank you to my sister Alanna whom I have leaned on for support throughout our youth and graduate studies and now with whom I compete for scholarships and publications!

To the most important person in my life, Sarah Sanderson, thank you for your endless patience, encouragement and love. Thank you for your understanding and inspiration and for supporting all of the decisions I have made as I pursue this academic career as a professional student. I would not be the person I have grown to be without you in my life.

Finally, I fully recognize that the mentorship of my supervisor Dr. Robert Tsushima since first attending one of his classes in late 2004 has afforded me the opportunity to develop into the person I am today. Robert, you have instilled in me the idea that in order to become a successful, independent researcher, one needs to look beyond the limitations of a specific field and embrace knowledge and scientific discovery in general. Thank you for the wonderful opportunity of pursing studies in your laboratory. You have immeasurably contributed to my development as an independent researcher. You have encouraged me to approach my project logically and critically, and have served as an ever-present source of insightful and thought-provoking discussion, support and counsel. Even more, thank you for your limitless friendship.

-iii- Table of Contents

Abstract ...... ii

Acknowledgments ...... iii

Table of Contents ...... iv

List of Abbreviations ...... vii

Chemicals & Compounds ...... ix

Symbols & Units ...... x

List of Tables ...... xi

List of Figures...... xii

Chapter 1: Introduction ...... 1

1.1 The ionic basis of cardiac contraction ...... 1

1.1.1 Ion channels & electrochemical gradients underlie cardiac contraction ...... 1 1.1.2 Cardiac action potential ...... 5 1.1.3 Cardiac channelopathies ...... 8

1.2 Potassium channels – structure & function ...... 14

1.2.1 Subunit assembly and stoichiometry ...... 15 1.2.2 Insights from atomic structures of potassium channels ...... 19

1.3 hERG Channels ...... 23

1.3.1 hERG channel structure ...... 23 1.3.2 hERG channel gating and kinetics ...... 25 1.3.3 Native hERG currents ...... 27 1.3.4 hERG channel pharmacology ...... 29 1.3.5 Posttranslational processing of hERG channels...... 31 1.3.6 Rescue of mutant hERG channels ...... 35

1.4 SNARE proteins ...... 37

1.4.1 Structure of SNARE proteins and mechanism of membrane fusion ...... 38 1.4.2 Modulation of ion channels by SNARE proteins ...... 41 1.4.3 Interaction of SNARE proteins and cardiac potassium channels ...... 44

Chapter 2: Research Objectives ...... 45

2.1 Rationale ...... 45

2.2 Hypothesis ...... 47

2.3 Specific aims and experimental design ...... 47 -iv- 2.4 Relevance ...... 48

Chapter 3: Materials and Methods ...... 50

3.1 DNA Constructs ...... 50

3.2 Generation of GST-fusion proteins ...... 50

3.3 Cell culture ...... 51

3.4 Transfection and drug treatment ...... 52

3.5 Electrophysiology ...... 54

3.6 Western blot analysis ...... 54

3.7 Antibodies ...... 57

3.8 Immunocytochemistry and confocal microscopy ...... 58

3.9 In vitro binding studies ...... 59

3.10 Coimmunoprecipitation ...... 59

3.11 Isolation of endogenous protein including membrane protein ...... 60

3.12 Immunoblot quantification and statistical analysis ...... 61

Chapter 4: Results ...... 63

4.1 Determining an ideal system for electrophysiological assessment of hERG channels ...... 63

4.2 STX1A significantly reduces hERG current amplitude ...... 67

4.3 STX1A affects steady-state inactivation but not gating kinetics ...... 68

4.4 STX1A-imparied hERG current amplitude is partially restored by E-4031 ...... 77

4.5 Colocalization of hERG and STX1A ...... 79

4.6 STX1A impairs hERG protein maturation ...... 80

4.7 Truncated hERG proteins as tools for the characterization of hERG-STX1A interactions ..... 89

4.8 hERG and STX1A binding experiments ...... 96

4.9 Endogenous expression of hERG & STX1A ...... 101

Chapter 5: Discussion ...... 104

5.1 hERG channel expression in HEK 293 cells is an appropriate model system ...... 105

5.2 STX1A-dependent reduction of hERG channel open probability ...... 106

-v- 5.3 STX1A-mediated reduction in PM expression of hERG channels ...... 111

5.4 STX1A-hERG binding experiments support functional data ...... 118

5.5 Disruption of STX1A-mediated impairment of hERG channel trafficking ...... 121

5.6 Endogenous expression and physiological relevance ...... 124

5.7 Conclusions ...... 127

5.8 Summary of recommendations for future experiments ...... 131

List of References ...... 133

Appendix 1 ...... 152

Preliminary results: Interaction of hERG and STX1A coexpression in tsA-201 cells ...... 152

Appendix 2 ...... 157

Preliminary results: Interaction of hERG and SNAP-25 coexpression in tsA-201 cells ...... 157

-vi- List of Abbreviations

ANOVA analysis of variance ANP atrial natriuretic peptide AP action potential APD action potential duration AV node atrioventricular node bpm beats per minute C closed ion channel state 2+ CaV voltage-gated Ca channel cDNA complementary deoxyribonucleic acid CF cystic fibrosis CFTR cystic fibrosis transmembrane conductance regulator cNBD cyclic nucleotide binding domain cRNA complementary ribonucleic acid DIC differential interference contrast DNA deoxyribonucleic acid EC50 effective drug concentration producing 50% of maximal response ECG electrocardiogram ECL enhance chemiluminescence EGFP enhanced green fluorescent protein ENaC amiloride-sensitive epithelial Na+ channel ER endoplasmic reticulum ERG ether-á-go-go-related gene FITC fluorescein isothiocyanate FKB 38 38-kDa FK506-binding protein, FKBP8 GI tract gastrointestinal tract GST glutathione S-transferase HA hemagglutinin Hc/sp 70 heat conjugate/stress-activated protein 70 HCN hyperpolarization-activated cyclic nucleotide-gated channel HEK 293 human embryonic kidney cell line 293 hERG human ether-á-go-go-related gene HRP horseradish peroxidase Hsp 90 heat shock protein 90 I inactivated ion channel state 2+ ICa inward Ca channel current 2+ ICa-L inward L-type Ca channel current If “funny” current IgG immunoglobulin G + IK1 inward rectifier K current + IKAch acetylcholine-activated inward rectifier K current + IKATP adenosine triphosphate sensitive inward rectifier K current + IKr rapid delayed rectifier K current + IKs slow delayed rectifier K current + IKur ultra-rapid delayed rectifier K current + INa inward Na channel current + 2+ INa/Ca Na / Ca exchanger + + INa/K Na /K ATPase current + Ito transient outward K current I-V current-voltage + + K2P four transmembrane, two pore K channel (background K channel)

-vii- Kir inward-rectifier K+ channel + KV voltage-gated K channel LQTS long QT syndrome M.W. molecular weight MiRP1 minK-related peptide 1 n sample size N.S. not significant + NaV voltage-gated Na channel NSF N-ethylmaleimide-sensitive factor O open ion channel state p probability PAS Per-Arnt-Sim PKA protein kinase A PM plasma membrane PVDF polyvinylidene fluoride rpm revolutions per minute S.E.M. standard error of the mean SA node sinoatrial node SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis SERCA sarcoplasmic/endoplasmic reticulum Ca-ATPase SM Sec1/Munc18-related proteins SNAP-23 synaptosome-associated protein of 23 kDa SNAP-25 synaptosome-associated protein of 25 kDa SNARE soluble N-ethylmaleimide-sensitive factor attachment protein receptor SR sarcoplasmic reticulum STX1 syntaxin 1 STX1A syntaxin 1A STX1B syntaxin 1B STX3 syntaxin 3 STX4 syntaxin 4 SUR sulfonylurea receptor synprint synaptic protein interaction site motif TdP torsade de pointes TM transmembrane TRITC tetra-methyl-rhodamine-isothiocyanate t-SNAREs target-membrane SNAREs V1/2 voltage required for half activation of current VAMP vesicle-associated membrane protein VAMP1 vesicle-associated membrane protein 1 VAMP2 vesicle-associated membrane protein 2 v-SNAREs vesicle-membrane SNAREs WT wild type

-viii- Chemicals & Compounds

Ar argon ATP (Mg salt) adenosine 5’-triphosphate magnesium salt ATP adenosine triphosphate BSA bovine serum albumin Ca2+ calcium CaCl2 calcium chloride cAMP cyclic adenosine monophosphate Cl- chloride CO2 carbon dioxide ddH2O double-distilled water DMEM Dulbecco’s modified eagle’s medium DTT dithiothreitol E-4031 1-[2-(6-methyl-2-pyridyl)ethyl]-4-(4-methylsulfonylaminobenzoyl)piperidine EDTA ethylenediaminetetraacetic acid EGTA ethylene glycol tetraacetic acid Endo H endoglycosidase H FBS fetal bovine serum glucose D-(+)-glucose (dextrose) H2O water HCl hydrochloric acid HeNe helium neon K+ potassium KCl potassium chloride KI potassium iodide KOH potassium hydroxide Mg magnesium MgCl2 magnesium chloride N2 nitrogen Na+ sodium NaCl sodium chloride NaOH sodium hydroxide NH4Cl ammonium chloride NP-40 nonidet P-40 O2 oxygen PBS phosphate buffered saline SDS sodium dodecyl sulfate TBS Tris-buffered saline TBST Tris-buffered saline with 0.1% tween 20

-ix- Symbols & Units

% percent ± plus or minus × g multiples of gravity ∆ deletion °C degrees Celsius Å angstrom cm centimeter d day g gram g/M grams per mole GΩ gigaohm h hour kDa kilodalton kg kilogram l liter M molar mg milligram min minute ml milliliter mm millimeter mM millimolar mmHg millimeters of mercury mV millivolt MΩ megaohm ng nanogram nm nanometer nm nanometer pA picoamp pF picofarad pH hydrogen ion concentration s second α alpha β beta μg microgram μl microliter π pi

-x- List of Tables

Table I: Long QT syndromes and known molecular correlates ...... 12

Table II: Cardiac potassium channel α-subunits and corresponding β-subunits ...... 18

Table III: Preparation of external solution for patch-clamp electrophysiology experiments ...... 56

Table IV: Preparation of internal solution for patch-clamp electrophysiology experiments ...... 56

Table V: Summary of syntaxin 1A interaction with K+ channels ...... 108

-xi- List of Figures

Figure 1. General anatomy of the heart and electrical conduction pathway ...... 2

Figure 2. Electrochemical gradients of major ion species present in cardiomyocytes ...... 4

Figure 3. Prototypical ventricular action potential and the associated ionic currents ...... 6

Figure 4. Long QT syndrome is caused by increased cardiac action potential duration ...... 9

Figure 5. General structure of K+ ion channel α-subunits ...... 16

Figure 6. K+ channel structural features ...... 20

Figure 7. hERG channel structure and gating ...... 24

Figure 8. LQT2 mutations cause a reduction in hERG channel whole cell current (IhERG) ...... 32

Figure 9. Structure and assembly of SNARE proteins ...... 40

Figure 10. Whole-cell mode of the patch clamp electrophysiology technique ...... 55

Figure 11. Expression of hERG WT cDNA in tsA-201 cells...... 64

Figure 12. Whole-cell currents elicited in stably transfected hERG-HEK 293 cells ...... 66

Figure 13. STX1A significantly impairs hERG current amplitude ...... 69

Figure 14. STX1A has no effect on hERG channel activation ...... 71

Figure 15. STX1A does not affect hERG channel deactivation ...... 73

Figure 16. STX1A does not affect fast inactivation or recovery from inactivation ...... 74

Figure 17. STX1A induces a hyperpolarizing shift in the midpoint of steady-state inactivation ...... 76

Figure 18. E-4031 can partially rescue STX1A-imparied hERG current amplitude ...... 78

Figure 19. Colocalization of hERG and STX1A ...... 81

Figure 20. Western blot analysis of hERG expression reveals two distinct bands ...... 83

Figure 21. STX1A reduces mature HA-hERG protein expression in a dose-dependent manner ...... 84

Figure 22. STX1A-mediated inhibition of HA-hERG channel maturation ...... 86 -xii-

Figure 23. Reduced temperature restores HA-hERG channel maturation ...... 88

Figure 24. Expression of hERG channel truncation mutations ...... 90

Figure 25. STX1A inhibits HA-hERG-Δ1120 maturation in a dose-dependent manner ...... 92

Figure 26. hERG and STX1A functionally interact downstream of residue 1000 ...... 93

Figure 27. E-4031 rescues STX1A-dependent inhibition of hERG-HA-Δ1045 maturation ...... 95

Figure 28. Interaction of hERG and STX1A ...... 97

Figure 29. hERG and STX1A interaction occurs between the residues 354 and 814 ...... 99

Figure 30. STX1A and hERG interaction is strongest with the shortest C-terminal truncations ...... 100

Figure 31. Endogenous expression of hERG and STX1A ...... 102

Figure 32. STX1A impairs hERG channel function - mechanism ...... 130

-xiii- -1-

Chapter 1: Introduction

1.1 The ionic basis of cardiac contraction

Vertebrate life would not be possible without the persistent, unrelenting beat of the heart, a muscular organ responsible for the pumping of blood throughout the body. In adult humans, the heart beats at an average rate of 72 bpm accumulating a total number of contractions of approximately three billion in an average lifetime (Marbán, 2002). The heart is such a vital organ, that it is the first functional organ system and begins pumping blood a mere 3 weeks after conception (Forouhar et al., 2006). The heart never rests; it relaxes so that blood can fill its atrial and ventricular chambers and then contracts so that blood can be forced throughout the body via the arteries. It is this cyclic repetition of relaxation and contraction that forms the basis of the heartbeat.

1.1.1 Ion channels & electrochemical gradients underlie cardiac contraction

Each heartbeat is initiated by a specialized group of pacemaker cells located in the sinoatrial (SA) node

(Fig. 1) (Baruscotti and Robinson, 2007; Maltsev and Lakatta, 2007). The SA node is said to have automaticity because it has an intrinsic ability for self-excitation. Modulation of heart rate can be affected via nervous or hormonal inputs as a result of exercise or emotional stimuli. The SA node, located in the upper wall of the right atrium, and subsequently the atrioventricular (AV) node, located in the lower right atrium; work in succession to control the rhythmicity of cardiac contraction. As an electrical impulse travels successively from the SA node to the AV node, atrial contraction occurs, forcing blood to flow into the ventricles. The electrical impulse is briefly delayed at the AV node, allowing for the ventricles to sufficiently fill before undergoing ventricular contraction. This occurs as the electrical impulse travels through the bundle of His and on to the Purkinje fibers. Propagation of electrical impulses through the myocardium is made possible by gap junction connections of neighboring myocytes and fibroblasts (Kohl et al., 2005).

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Figure 1. General anatomy of the heart and electrical conduction pathway

Initiation of electrical excitability in the heart occurs spontaneously in the SA node (blue) which sends electrical impulses throughout the myocardium via the AV node. This initiates atrial systole (contraction), the first step of the cardiac cycle. After a short delay, the electrical impulse continues travelling from AV node through the bundle of His (red), and onto the Purkinje fibers which causes ventricular systole. The final step in the cardiac cycle is complete cardiac diastole (relaxation).

Figure adapted from Anatomy of the Human Body, 20th edition, Gray, H. (1918) pp 507

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Heart cells, or myocytes, are characterized by carefully maintained internal ionic gradients, which produce an electrochemical gradient (Fig. 2). A sophisticated selectively permeable plasma membrane (PM) surrounding each cardiomyocyte allows for the precise coordination of ion-specific transmembrane proteins called channels and pumps (Gouaux and Mackinnon, 2005). Ion channels have the remarkable ability of permitting flow of molecules through their highly selective pores at rates exceeding 106 molecules per s (MacKinnon, 2004). In contrast, transporters and pumps operate at much slower rates, and are fueled by electrochemical gradients or ATP, which cause conformational changes in these membrane-spanning proteins.

Cardiomyocytes are partially permeable to K+ ions while at rest, and the outward conductance of these ions produce a negative electrochemical gradient with respect to the extracellular space, resulting in a resting membrane potential of approximately -80 mV. During excitation, the membrane potential reverses as a result of the opening of Na+-selective channels. This process is referred to as cardiac depolarization. Depolarization and the inward conductance of Na+ ions leads to the opening of

2+ 2+ voltage gated Ca channels (CaV) and the influx of Ca ions down their electrochemical gradient. An increase in intracellular Ca2+ triggers the opening of ryanodine receptors connected to the sarcoplasmic reticulum (SR). This Ca2+-induced Ca2+-release greatly increases the intracellular concentration of that ion, in turn causing activation of the cardiac contractile machinery. Cardiac relaxation is achieved following the closure of Na+ and Ca2+ channels. After a delay, K+-selective ion channels open, thereby defining the absolute refractory period and restoring the negative electrochemical gradient in preparation for the initiation of another cardiac cycle. This process is called repolarization. Several types of K+-selective ion channels contribute to cardiac repolarization, each with unique structure determining temporal and voltage-dependent current characteristics

(Nerbonne, 2000). In fact, the expression of cardiac ion channels is not homogeneous throughout the heart, thereby allowing for the localized expression of ion channels defining the highly tuned and temporally dependent nature of the cardiac cycle.

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Figure 2. Electrochemical gradients of major ion species present in cardiomyocytes

Schematic representation of four types of ions and generalized voltage-gated ion channels through which they conduct. The extracellular [Na+] (purple) and [Ca2+] (orange) are higher than inside the cardiomyocyte and the major conductivity of these ion species occurs during depolarization. [Cl-] is higher outside of the myocyte and conductance occurs in both directions. Repolarization of the myocardium is driven by the movement of K+ ions (green) down their concentration gradient across the plasma membrane of the cardiomyocyte. Several different K+ ion channels underlie cardiac repolarization.

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1.1.2 Cardiac action potential

The cardiac action potential represents the summation of the electrical activity of ion channels and transporters in a particular myocyte during the cardiac cycle. Fig. 3 illustrates a representative cardiac action potential for prototypical ventricular myocyte (Sanguinetti and Tristani-Firouzi, 2006). The individual depolarizing and repolarizing currents are defined below with the probable clone and nomenclature of the genes encoding the proteins listed next to the representative currents. The cardiac action potential is defined by five distinct phases.

Phase 0 is defined by the upstroke of the action potential. Several independent ionic currents are responsible for the electrical impulse producing cardiac depolarization; however, voltage-gated Na+ channels (INa) are primarily responsible for the initiation of the cardiac action potential. NaV1.5 channels are encoded by the SCN5A gene which open (activate) rapidly, and depolarize the myocyte membrane potential to greater than +40 mV (Catterall, 1996). Despite maintained depolarization, Na+ channels rapidly close, a process called inactivation, and they are very unlikely to open again during the remainder of the cardiac action potential.

Phase 1 is defined by an initial repolarization of the action potential, and the appearance of a distinctive “notch” immediately following phase 0. K+ currents are in the outward direction promoting repolarization, and operate under strict voltage and temporal conditions. The transient outward current (Ito) is responsible for early repolarization and is comprised of two components. Ito1 is

+ predominated by the voltage-gated K channel KV4.2/4.3 encoded by the KCND gene which produces this fast activating and inactivating channel (Birnbaum et al., 2004). Additionally, Ito2 has been shown to be sensitive to intracellular changes in Ca2+, but the identity of the molecular correlate is unknown.

+ The voltage-gated KV1.5 channel underlies the ultra-rapidly activating delayed K current (IKur) and is encoded by the KCNA5 gene (Tamargo et al., 2004).

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Figure 3. Prototypical ventricular action potential and the associated ionic currents

A) Generalized representation of the predominant ion channels involved in ventricular depolarization and repolarization, and the Na+/Ca2+ exchanger. B) Typical electrophysiological recording from a ventricular myocyte illustrating a cardiac action potential. There are 5 distinct phases, with phase 0, upstroke of the action potential, representing depolarization. C) Representative current traces from the various ion channels and transporter underlying the cardiac action potential. Depolarizing currents are provided by Na+ (purple) and Ca2+ (orange) ion channels, whereas repolarizing currents are produced by K+ ion channels (green). Corresponding genes and molecular correlates are indicated to the right of the current traces.

Figure adapted with permission from Macmillan Publishers Ltd: Nature (Marban, 2002), copyright (2002).

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Phase 2 represents the plateau phase of the cardiac action potential. This phase involves a balance in

2+ + 2+ the depolarizing Ca current (ICa-L) and the various repolarizing K currents. L-type Ca channels

(CaV1.2) are responsible for ICa-L current, and they are encoded by the CACNA1C gene. These channels open rapidly following depolarization, however, unlike INa current, they inactivate slowly and not completely, contributing considerably to the current plateau (Marbán, 2002). Simultaneously, several voltage-dependent K+ channels underlie the repolarizing K+ currents. These channels have reduced conductances at positive transmembrane potentials, thereby also prolonging the plateau phase. This is important to insure an adequate supply of intracellular Ca2+ for contraction, as well as for establishing an absolute refractory period to prevent the generation of a re-entrant arrhythmia. The

+ rapidly- and slowly-activating delayed rectifier K currents (IKr and IKs) activate substantially more slowly than IKur, and contribute to the later part of phase 2 repolarization (Sanguinetti and Tristani-

Firouzi, 2006). IKr currents are produced by KV11.1, the human ether-á-go-go-related gene (hERG) channel which is encoded by the gene KCNH2 and is discussed in more detail below. IKs currents are produced by KV7.1 channels (KvLQT), which are encoded by the gene KCNQ1.

Phase 3 of the cardiac action potential is characterized by repolarization due to the reduction of ICa-L

+ and an increase in the K currents IKr and IKs. IKr is the most important component of phase 3 repolarization as its conductance increases as the membrane potential becomes progressively more

+ negative. Additionally, IK1, an inward rectifying K current produced by the Kir2.1 channel and encoded by the KCNJ2 gene opens as the resting membrane potential is restored (Lehnart et al., 2007).

Phase 4 simply represents the resting membrane potential. Normally, the resting membrane potential of cardiac myocytes is roughly -80 mV. This extremely negative membrane potential is the result of the PM being substantially more permeable to K+ ions than any other ion species, thereby driving the membrane potential toward the K+ equilibrium potential. A slew of ion pumps and exchangers aid in the maintenance of the electrochemical gradient, however, it is primarily controlled by IK1. The

+ 2+ 2+ Na /Ca exchanger (INa/Ca) is encoded by the NCX1 gene and allows for the removal of 1 Ca ion for 3

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Na+ ions (Philipson and Nicoll, 2000). Na+ ions flow down their electrochemical gradient across the PM driving the counter-transportation of Ca2+. Active transport of Na+ and K+ ions against their concentration gradients helps to maintain the electrochemical gradients (INa/K). The gene ATP1A encodes the energetically demanding Na+/K+ATPase. One ATP molecule is hydrolyzed in order to pump 3 Na+ ions out of the cell while 2 K+ ions are pumped into the cell, both ion species against their concentration gradient. Finally, cells of the SA-node also possess a pacemaker current or “funny current” (If), which is produced by the hyperpolarization-activated, cyclic nucleotide-gated channel

(i.e. HCN2). Interestingly, this poorly selective cation channel activates following hyperpolarization, allowing the mixed Na+/K+ inward current underlying the automaticity of the SA node (Accili et al.,

2002).

Together, the myriad of ion channels and transporters working under specific temporal- and voltage- dependent constraints produce the cardiac action potential, which provides the basis for the cardiac cycle. However, as mentioned above, the composition of ion channels throughout the myocardium is not consistent. Regional differences in expression and function of these channels allows for the choreographed conduction of an electrical impulse originating at the SA node and propagating throughout the heart, producing the carefully timed sequence of atrial and ventricular contractions of the cardiac cycle (Nerbonne, 2000).

1.1.3 Cardiac channelopathies

The electrocardiogram (ECG) is a tool used to assess normal cardiac function and represents an averaged electrical gradient generated by cardiomyocytes plotted versus time. Fig. 4 illustrates a typical ECG whose distinct waveform pattern corresponds to the temporally complex sequence of atrial and ventricular depolarization and repolarization (Sanguinetti and Tristani-Firouzi, 2006). The

ECG is obtained using surface electrodes placed on the body. The initial P-wave corresponds to atrial activity, and the QRS complex represents ventricular depolarization, particularly the initial upstroke of

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Figure 4. Long QT syndrome is caused by increased cardiac action potential duration

A) Representative cardiac action potential traces from normal (left) and LQTS (right) ventricular myocytes. Action potential duration is longer in LQTS myocytes. B) Representative ECG traces from normal (left) and LQTS (right) subjects. The ECG waveform is characterized by a P-wave corresponding to atrial activity, the QRS complex corresponding to ventricular depolarization, and a gently rolling T-wave, corresponding to ventricular repolarization. The QT interval is prolonged in the LQTS trace, which increases the likelihood of early afterdepolarizations and ventricular tachycardia. C) ECG trace representing a torsade de points arrhythmia.

Figure adapted with permission from Macmillan Publishers Ltd: Nature (Sanguinetti and Tristani-Firouzi, 2006), copyright (2006)

-10- the cardiac action potential. The elongated T-wave represents ventricular repolarization. The QT- interval is the period of time from the beginning of the QRS complex to the end of the T-wave and is a measurement used to assess the length of time required for ventricular repolarization during a single cardiac cycle. Normal cardiac rhythm can be assessed by the ECG as can the presentation of abnormal cardiac arrhythmias. Early afterdepolarizations, for example, occur when a region of the heart begins another cycle of depolarization before repolarization has completed (Knollmann and Roden, 2008).

Cardiomyocytes are electrically coupled to one another, which allows for the propagation of rogue electrical impulses interrupting the normal rhythm of cardiac repolarization. The production of self- perpetuating “wavelets” of electrical activity from an early afterdepolarization can produce ventricular fibrillation or tachycardia, recorded as a series of long or short wavelets, respectively, on the ECG

(Marbán, 2002). Ventricular tachycardia produces an uncoordinated series of fast irregular heartbeats, which can degenerate into torsade de pointes (TdP) arrhythmia, characterized by a twisting of the ECG around its isoelectric axis (Keating and Sanguinetti, 2001). TdP can revert to normal sinus rhythm, or can degenerate into ventricular fibrillation. This type of arrhythmia can cause syncope and death if cardiopulmonary resuscitation and defibrillation is not performed within minutes of onset.

Because of the complex nature and composition of the ion channels underlying cardiac repolarization, slight perturbations in the function of a small number of channel proteins can have compounding effects on overall cardiac electrical conduction (Marbán, 2002). Channelopathies refer to mutations of ion channels which are linked to inherited diseases (Ashcroft, 2006). Such mutations can have loss-of- function or gain-of-function effects. Long QT syndrome (LQTS) is defined by a prolongation of the QT interval, greatly increasing the risk of ventricular fibrillation and TdP. In addition to acquired forms of

LQTS, the disease can be the result of congenital channelopathies affecting cardiac ion channel function or mutations in proteins associated with ion channels, leading to perturbations in the normal cardiac rhythm (Subbiah et al., 2004). Once considered a fairly rare disease, LQTS has recently been hypothesized to affect more than 1 in 2,500 individuals (Crotti et al., 2008). LQT-related episodes resulting in disrupted cardiac rhythm may be precipitated by extreme physical or emotional stress.

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Currently, there are 10 classes of LQTS which are outlined in Table I (Sanguinetti and Tristani-Firouzi,

2006; Wolf and Berul, 2006). LQTS requires inheriting only one variant of these gene products, however, more genes associated with LQTS remain to be discovered as approximately 30-35% of patients cannot be linked to any of the known genes (Schwartz, 2005; Crotti et al., 2008). Dominant- negative congenital mutations in either KCNQ1 (KvLQT1) or KCNH2 (hERG), the α-subunits underlying

IKs or IKr are the most common cause of LQTS (Marbán, 2002). Reductions in IKs or IKr result in a prolonged QT interval, on the order of 2-5% due to increases in cardiac action potential duration

(APD). Point mutations in KvLQT1 cause LQT1, the most common form of congenital LQT accounting for 30-35 % of all cases (Lehnart et al., 2007). There are two variants of LQT1 – an autosomal dominant form called Romano-Ward syndrome, and a much rarer recessive form called Jervell and Lange-

Nielsen syndrome which is defined by profound sensorineural deafness in addition to LQTS (Wang, Q. et al., 1996).

Deletions, missense mutations and splice-donor mutations in hERG channels cause LQT2 which accounts for 25-30 % of all LQTS cases (Curran et al., 1995; Lehnart et al., 2007). To date, 291 hERG channel mutations have been identified (see http://www.fsm.it/cardmoc/). Generally, many LQT2 mutations result in disrupted hERG channel folding and trafficking to the PM, resulting in greatly reduced or completely abolished current amplitude (Anderson et al., 2006). Most acquired forms of

LQTS are the result of drug action which blocks highly sensitive hERG channels. Unique properties of the hERG channel pore structure make it a vulnerable target to unintentional drug interaction. For this reason, all drugs developed must be screened for hERG channel blockade. hERG channels are highly prone to block by a wide variety of drug types including antihistamines and antibiotics (Tseng, 2001;

Vandenberg, J. I. et al., 2001). Ironically, the hERG-channel blockers quinidine and dofetilide were used in the early treatment of arrhythmias before being withdrawn by regulatory agencies.

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Table I: Long QT syndromes and known molecular correlates

Loci Gene Protein Symptoms Current Effect

LQT1 11p15.5 KCNQ1 KvLQT1 Romano-Ward and IKs Loss-of-function (KV7.1) Jervell and Lange- (reduced current) Nielsen (autosomal recessive) LQTS

LQT2 7q35-36 KCNH2 hERG Romano-Ward LQTS IKr Loss-of-function (KV11.1) (reduced current)

LQT3 3p21-24 SCN5A NaV1.5 Romano-Ward LQTS INa Gain-of-function (impaired inactivation)

LQT4 4q25-27 ANK2 Ankyrin B Romano-Ward LQTS INa/Ca Loss-of-function INa/K (Impairs trafficking of channels affecting resting membrane potential)

LQT5 21q22 KCNE1 MinK Romano-Ward and IKs Loss-of-function Jervell and Lange- (reduced current) Nielsen (autosomal recessive) LQTS

LQT6 21q22.1 KCNE2 MiRP1 Romano-Ward LQTS IKr Loss-of-function (reduced current)

LQT7 17q23 KCNJ2 Kir2.1 Anderson-Tawil IK1 Loss-of-function syndrome (reduced current)

LQT8 12p13.3 CACNA1C CaV1.2 Timothy syndrome ICa-L Gain-of-function (impaired inactivation)

LQT9 3p25.3 CAV3 Caveolin 3 Romano-Ward LQTS INa Gain-of-function (impaired inactivation)

LQT10 11q23.3 SCN4B NaVβ4 Romano-Ward LQTS INa Gain-of-function (impaired inactivation)

-13-

Mutations affecting cardiac Na+ channel inactivation define LQT3 (Wang, Q. et al., 1995). Unlike mutations affecting cardiac K+ channels, LQT3 is the result of a gain-of-function mutation which reduces inactivation in NaV1.5, thereby increasing the depolarizing inward current during the plateau phase and increasing APD (Remme et al., 2008). Lidocaine and mexiletine are anesthetic agents which specifically block the non-inactivating component of LQT3 and may be a useful treatment option.

Ankyrin-B is an intracellular adaptor protein encoded by the gene ANK2 that is expressed throughout the myocardium and is important in protein trafficking. LQT4 caused by mutant Ankyrin-B results in a loss of function of that protein which leads to decreases in the expression of the IP3 receptor, the

Na+/Ca2+ exchanger and the Na+/K+ ATPase, all of which are essential for the maintenance of resting membrane potential and intracellular [Ca2+] (Schott et al., 1995; Mohler et al., 2003). This leaves the myocardium susceptible to early after and delayed depolarizations.

In addition to mutations in the α-subunits encoding the cardiac ion channels, mutations in the β- subunits which coassemble with α-subunits to form function channels also cause LQTS. Although less common, mutations in KCNE1 (LQT5) and KCNE2 (LQT6) slow action potential repolarization, thereby increasing APD (Splawski et al., 1997; Abbott et al., 1999). KCNE1 is the gene which encodes MinK, an accessory β-subunit to KvLQT1 and a required component of IKs. In rare homozygous forms it can cause Jervell and Lange-Nielsen syndrome. MinK-related peptide 1 (MiRP1) is encoded by KCNE2 which encodes the β-subunit postulated to be involved with the IKr current. Mutations in these β- subunits may cause alterations in counterpart α-subunit expression, may cause the formation of nonfunctional protein, or act in a dominant negative manner to suppress current.

Anderson-Tawil syndrome (LQT7) is characterized by periodic paralysis of skeletal muscles, LQTS and skeletal deformities (Tristani-Firouzi et al., 2002). It is caused by a mutation in Kir2.1 encoded by the gene KCNJ2. Although symptoms are highly variable between individuals, mutations in Kir2.1 have been found to act in a dominant-negative manner, suppressing IK1 current and causing a delay in the

-14- final stage of ventricular repolarization. Timothy syndrome is another rare LQTS (LQT8) involving a mutant ion channel. Mutations in CACNA1C, the gene encoding the α-subunit of the L-type Ca2+ channel (CaV1.2) result in a severe prolongation of the cardiac action potential (Splawski et al., 2004).

In addition, the disorder afflicts multiple organ systems and symptoms include structural abnormalities of the heart, syndactyly (partial fusion of fingers and toes), immune deficiency and autism. Two mutations have been characterized in highly conserved regions of the Ca2+ channel affecting inactivation, producing a gain-of-function effect and thereby increasing APD.

Finally, LQT9 and LQT10 have been characterized and shown to involve mutations in proteins affecting INa inactivation. LQT9 is caused by a mutation in CAV3 which encodes the NaV1.5-interacting protein caveolin 3 (Vatta et al., 2006). LQT10 is caused by a mutant SCN4B encoding a β-subunit for INa

– NaVβ4 (Van Norstrand et al., 2007). Both of these LQT syndromes involve gain-of-function mutations.

Interestingly, in addition to loss-of-function mutations causing LQTS, a few gain-of-function mutations exist for cardiac ion channels involved in cardiac repolarization leading to a shortened Q-T interval.

These syndromes are termed Short QT syndrome. Several examples of such gain-of-function mutations have been characterized for IKs (KvLQT1), IKr (hERG) and IK1 (Kir2.1), most of which abolishing or impairing channel inactivation, or preventing or delaying channels from closing (Brugada et al.,

2005). The vast array and variations of cardiac channelopathies attest to the complex nature of cardiac electrophysiology.

1.2 Potassium channels – structure & function

Potassium channels are expressed in every living organism, and are probably the oldest group of ion channels. In humans alone, over 80 K+ channel genes have been characterized (Coetzee et al., 1999;

Roden et al., 2002). Although there is great diversity and variety in the types of K+ ion channels, they are all related members of a single protein family, and can be easily recognized by a highly conserved

-15- amino acid sequence called the K+ channel signature sequence (MacKinnon, 2003). K+ channels are α- helical transmembrane (TM) proteins which are able to selectively permit the flow of K+ ions at rates approaching the diffusion limit, into the cell down their electrochemical gradient. The regulation of K+ ion conduction is important for numerous physiological processes, including the regulation of cell volume, hormonal secretion, and the production of electrical impulses as in the excitable myocardium.

1.2.1 Subunit assembly and stoichiometry

Three common groups of K+ channel families include the inward rectifiers, background channels, and voltage-gated channels (Fig. 5). Although these channels are structurally diverse, all possess the highly conserved K+ channel signature sequence, which forms a structural element called the selectivity filter (Heginbotham et al., 1994). The selectivity filter lies in the pore region of the channel, through which K+ ions flow, and is surrounded by four usually identical subunits that are symmetrically arranged around an ion conduction pathway (Doyle et al., 1998).

Inward rectifiers are the simplest group of K+ channels. They are characterized by permitting the conductance of current in the inward direction and open at negative membrane potentials where the electrical force of K+ ions overcomes the concentration gradient. The pore-forming α-subunits of inward rectifiers are composed of two TM domains connected by the pore region, and containing both intracellular N- and C-termini. Four such monomers amalgamate to form a functional channel, which is said to have tetrameric structure. The inward-rectifier K+ channels are noted Kir(x) where x represents channels of the same subfamily. Members of each subfamily are able to coassemble to form a functional channel. For example, combinations of four Kir2.1 and Kir2.2 monomers, members of the same Kir2 subfamily, are capable of forming heteromeric channels with one another, or can form homomeric channels alone.

-16-

Figure 5. General structure of K+ ion channel α-subunits

Three families of K+ channels are illustrated. A) Inward rectifier K+ channels are composed of 2 TM α-helices and 1 pore region. Four such α-subunits are required to form a functional tetrameric channel. B) Background K+ channels, also called “leak channels”, are composed of 2 pore regions and 4 TM domains. Two such α-subunits are required to form a functional dimeric channel. C) General schematic for a voltage-gated K+ channel α- subunit. These tetrameric channels require 4 α-subunits composed of 6 TM domains and 1 pore region each. TM domains are noted S1-S6, with the S4 domain containing 4 highly conserved, positively charger Arg residues. N- and C-terminal domains for all K+ channel families are located on the cytoplasmic side of the PM.

-17-

Several inward rectifier K+ channels are expressed in the heart (Table II). These channels underlie the

IK1, IKAch and IKATP currents which are essential for the maintenance of the resting membrane potential, hyperpolarizing depolarized membranes, and contributing to repolarization (Roden et al., 2002). IK1 is encoded by Kir2.1 and Kir2.2, and plays an important role in terminal repolarization as well as maintenance of the resting membrane potential. Inward rectifier channels can open in response to

+ ligand binding. The acetylcholine-gated inward rectifier K channel current (IKAch) requires the coexpression of both Kir3.1 and Kir3.4, and opens in response to the binding of G-protein subunits.

This current is highly expressed in atrial myocytes, helping to reduce heart rate following stimulation of the vagus nerve (Nishida et al., 2007). The adenosine triphosphate (ATP)-sensitive inward rectifier K+ channel current (IKATP) is encoded by 4 Kir6.2 α-subunits and 4 sulfonylurea receptor 2A (SUR2A) β- subunits. SUR2A is composed of 12 TM domains and 2 ATP-binding cassettes. Kir6.2/SUR2A channels open in response to a decrease in the concentration of intracellular ATP, thus linking the metabolic state of the cell to electrophysiological activity.

+ Background K channel α-subunits, also known as “leak channels” and noted K2P(x), are composed of 4

TM domains and 2 pore regions with intracellular N- and C-termini (Birnbaum et al., 2004). In order to form a functional dimeric channel, two such α-subunits are required. Channels can be homomeric or heteromeric. K2P currents are not time-dependent and show little voltage-dependence (Roden et al.,

2002). The four classes of K2P channels expressed in the heart include TASK, TWIK, TREK and THIK, which are believed to be important for maintenance of the resting membrane potential and myocyte excitability. Several factors are known to regulate these channels including oxygen tension, pH, mechanical stretch, and G-proteins. Generally, these channels operate as inward rectifier channels and the 9 subfamilies known to be expressed in the human heart are listed in Table II.

+ Voltage-gated K channel (KV) α-subunits consist of 6 TM domains (S1-S6) with a pore loop located between S5 and S6 and possess intracellular N- and C-terminal domains. The pore region, consisting of the pore loop and S5 and S6 TM segments are structurally similar to Kir channel subunits. KV

-18-

Table II: Cardiac potassium channel α-subunits and corresponding β-subunits

α-Subunit β-Subunit

Current Name Gene Name Gene

Inward-rectifier K+ channels

IK1 Kir2.1 (IRK1) KCNJ2 Kir2.2 (IRK2) KCNJ12

IKAch Kir3.1 (GIRK1) KCNJ3 Kir3.4 (GIRK4) KCNJ5

IKATP Kir6.2 (BIR) KCNJ11 SUR2A ABCC9

Background K+ channels

K2P K2P1.1 (TWIK-1) KCNK1

K2P2.1 (TREK-1) KCNK2

K2P3.1 (TASK-1) KCNK3

K2P5.1 (TASK-2) KCNK5

K2P6.1 (TWIK-2) KCNK6

K2P9.1 (TASK-3) KCNK9

K2P10.1 (TREK-2) KCNK10

K2P13.1 (THIK-1) KCNK13

K2P17.1 (TASK-4) KCNK17

Voltage-gated K+ channels

IKs KV7.1 (KvLQT1) KCNQ1 minK KCNE1

IKr KV11.1 (hERG) KCNH2 minK KCNE2 MiRP1 KCNE2

IKur KV1.5 (HK2) KCNA5 KVβ1 (KVβ3) KCNAB1

KVβ2 KCNAB2

Ito1 KV4.3 KCND3 KChIP2 KCNIP2

KV1.4 KCNA4

KV4.1 KCND1 KChIP1 KCNIP1

KV4.2 KCND2 KChIP2 KCNIP2

-19- channel monomers differ such that they possess 4 additional TM domains, including the S4 segment which contains positively charged residues critical for sensing changes in the transmembrane potential. KV channels are tetrameric in structure and can be homomeric or heteromeric

(combinations of different α-subunits from the same subfamily). There are 4 voltage-gated K+ currents found in the heart: IKr, IKs, IKur, and Ito1. The channels underlying these currents were discussed above, and are also listed with their known β-subunits in Table II.

1.2.2 Insights from atomic structures of potassium channels

X-ray crystallographic studies from the MacKinnon laboratory at the Rockefeller University have revealed the atomic structures of several K+ channels in the last decade. The crystal structure of KcsA, a bacterial K+ channel containing 2 TM domains and a pore domain in each α-subunit, is homologous to all K+ channels, and revealed that the pore structure resembles an inverted teepee, with an ion selectivity filter located near the extracellular end (Fig. 6) (Doyle et al., 1998). Lessons learned from this atomic structure provided a glimpse at the complex inner workings of K+ channels, finally allowing investigators to correlate mechanistic theory of ion channel function with intricate structural detail.

Recently, crystal structures have been obtained for several ion channels and ion channel domains

+ + including: KvAP, a voltage-dependent bacterial K channel; MthK, a ligand-gated K channel; Kv1.2

(Shaker), a mammalian voltage-dependent K+ channel; and most recently a Kir3.1-prokaryotic Kir channel chimera (Jiang et al., 2002; Jiang et al., 2003a; Long et al., 2005b; Nishida et al., 2007). Here I will briefly discuss observations obtained from these crystallographic structures related to K+ channel selectivity, gating, and voltage-sensing.

K+ channels are able to select over Na+ ions by a factor of more than 103, despite the fact that the atomic radius of K+ is 1.33 Å, substantially larger than the atomic radius of Na+. Perhaps most impressively, selectivity occurs as ions travel through channels at rates approaching the diffusion limit.

This is achieved by the specific architecture of the pore region formed by the four α-subunits which

-20-

Figure 6. K+ channel structural features

A) The KcsA K+ channel was crystallized in the closed conformation. 2 α-subunits are shown revealing the ion conduction pathway and the selectivity filter occupied by 2 K+ ions (white spheres). Gly (red) and Tyr (yellow) residues are also indicated. B) Two KV1.2 (Shaker) α-subunits are shown in the open conformation. Only 2 TM domains (S5 and S6) and the pore region are illustrated for simplicity. S6 hinges at a Gly residue, providing + access for the flow of K ions from the cytoplasm. C) A single KV1.2 α-subunit with all 6 TM domains. The entire

KV1.2 channel with a single α-subunit viewed from the side (D) and from below (E). S1-S4 domains (voltage sensor) are symmetrically distributed in units around the channel pore.

Figure adapted with permission from Macmillan Publishers Ltd: Nature (Sanguinetti and Tristani-Firouzi, 2006), copyright (2006).

-21- make up the channel. Near the midpoint of the PM, the diameter of the ion conduction pathway, formed by the inner α-helices of each subunit, is 10 Å. This water-filled vestibule allows K+ ions to remain hydrated until immediately before passing through the selectivity filter. This serves to lower the electrostatic repulsive forces intrinsic to the hydrophobic membrane interior. Additionally, tilted pore α-helices, passing only partially through the PM are negatively charged and pointed toward the ion conduction pathway. The selectivity filter possesses a highly conserved sequence of amino acid residues: Thr-Val-Gly-Tyr-Gly (the K+ channel signature sequence). The side-chains of these amino acids, consisting of one hydroxyl group and four carbonyl oxygen atoms on each subunit, face towards the narrow ion-conduction pathway and form the basis for four evenly spaced octahedral K+ ion binding sites. Dehydrated K+ ions are perfectly stabilized by four upper and four lower oxygen atoms, thereby mimicking the K+ ion hydration shell. The selectivity filter accommodates 2 K+ ions, either in the 1,3 or 2,4 configuration, with ions separated by a single water molecule (Doyle et al.,

1998). These positions ensure that repulsive forces between K+ ions drive rapid conduction down the electrochemical gradient through the selectivity filter while preventing the ions from binding with the selectivity filter itself.

The crystallographic structures of the first two K+ channels provided evidence for a generalized mechanism of K+ channel gating. KcsA was crystallized in the closed conformation, while MthK was in the open conformation. The KcsA channel was similar in structure to MthK, with the exception of straight inner pore α-helices which formed a tight bundle near the intracellular membrane. This bundle had an opening of 3.5 Å and was lined with hydrophobic residues, thereby providing a barrier to K+ ion access. Alternatively, the inner α-helices of MthK were bent at a hinge point located in the

PM below the selectivity filter, splaying the helices such that the central cavity of the channel pore becomes accessible from the cytoplasm. Furthermore, this Gly residue hinge point is highly conserved among K+ channels. Thus, large conformational changes within the transmembrane domains underlie pore opening.

-22-

Voltage-gated K+ channels possess 6 TM domains, a pore region and are tetrameric in structure. S5 and S6 form the central pore between the subunits, and S1-S4 form the voltage sensors symmetrically distributed around the pore region. Crystal structures from the MacKinnon lab of KvAP and KV1.2 have shed some light on the mechanism by which KV channels sense changes in the membrane potential and translate them into conformational changes of the pore structure. KV channels are able to open and close in response to changes in transmembrane potential, thereby providing a feedback mechanism for the operation. The KvAP atomic structure revealed the importance of the S4 domain which possesses four positively charged Arg residues per subunit. These charged amino acid residues represent gating charges which travel through the PM in response to changes in the membrane potential, thereby coupling electrical work to channel opening. MacKinnon’s group has gone on to assert that these gating charges, located on hydrophobic helix-turn-helix structures, resemble voltage sensor paddles which operate as independent units travelling a distance of 15-20 Å across the PM during channel activation (Jiang et al., 2003b; Ruta et al., 2005). Critics of the paddle theory assert that it is naïve to think of the voltage sensors as “buoys” floating in the plasma membrane and suggest that the coupling of voltage sensor movement to changes in pore conformation is more complex and dynamic, requiring further characterization and study (Roux, 2006). Alternatively, evidence from biophysical analysis of gating charge movement suggests that the voltage sensor is only required to move 5 Å (Laine et al., 2004; Posson et al., 2005). Critics argue that MacKinnon’s group cannot make a definitive conclusion regarding voltage sensor movement without the crystal structure of a KV channel in the closed state. That said, it is undeniable that work done in this field over the last decade has advanced our understanding of ion channel structure and function exponentially, helping us to comprehend the mechanisms underlying the most basic and essential physiological processes in nature.

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1.3 hERG Channels

+ The pore-forming α-subunit underlying the rapidly activating delayed rectifier K current (IKr) in the heart is encoded by KCNH2, the human ether-á-go-go-related gene (hERG) channel. In addition to the plethora of congenital mutations associated with this channel causing LQT2, a wide variety of compounds block hERG currents, causing the acquired form of the syndrome. In this section, I will discuss hERG channel structure and function, trafficking and expression, and pharmacology.

1.3.1 hERG channel structure

Much of what is known about hERG channel structure is derived from studies of homologous K+ channels including KcsA and KV1.2 (Doyle et al., 1998; Long et al., 2005b). Similar to KV1.2, hERG channels are tetrameric in structure, composed of four identical α-subunits each consisting of 6 TM domains (S1-S6) and a pore region (Fig. 7 A) (Roden and George, 1997). Functionally, each subunit can be divided into 2 main components: a K+-selective pore (S5-S6) and a transmembrane voltage sensor

+ (S1-S4). The K selectivity filter for hERG differs slightly from other KV channels and is composed of the residues Ser-Val-Gly-Phe-Gly, however, it is believed that the structure of the selectivity filter is not affected. Additionally, hERG channels differ from typical KV channels in that they possess elongated α- helical pore loops located between S5 and S6. These pore loops lack the hydrogen bonds present in other KV channels which help to stabilize the channel in the open conformation. The presence of larger, flexible S5-P loop linkers in the outer mouth of the channel are postulated to affect Na+/K+ selectivity, as well as hERG’s rapid voltage-sensitive inactivation process. hERG channel activation, as in other KV channels, is made permissible by a hinging of S6 α-helices, allowing for cytoplasmic access to the ion conduction pore. Located two helical turns below the typical Gly residue, hERG features an

Ile-Phe-Gly motif, which serves as an activation hinge point without permitting channel closing (Long et al., 2005b). For this reason, upon depolarization of membrane potential, hERG channels slowly activate and then rapidly inactivate without closing. Repolarization of the membrane potential causes

-24-

Figure 7. hERG channel structure and gating

A) Diagram of a single hERG channel α-subunit. hERG channels are characterized by an unusually long S5-P linker and the presence of an N-terminal PAS domain and a C-terminal cNBD domain. Four identical subunits are required to assemble a functional hERG channel. B) Simplified gating schematic for hERG channels. Upon depolarization, hERG channels activate with slow kinetics, and then inactivate with fast kinetics. Channels cannot close from the inactivated state. Following repolarization, hERG channels recover from inactivation with fast kinetics and momentarily transition through the open state, producing a distinctive tail current. hERG channels deactivate with slow kinetics upon further repolarization.

-25- hERG channels to rapidly recover from inactivation, opening momentarily, and then slowly deactivating and returning to the closed conformation. Thus, unique structural properties endow hERG channels with a distinctive gating mechanism (Fig. 7 B).

hERG channel α-subunits also possess large intracellular N- and C-terminal domains. The 135 amino acid Per-Arnt-Sim (PAS) domain, located at the N-terminus, is a structure found throughout the phylogeny of nature. First discovered in the Drosophila proteins PER and ARNT, the PAS domain is important for protein-protein interactions that are involved in environmental sensing and transcriptional regulation (Huang et al., 1993). Currently, it is unknown whether the PAS domain serves a similar function in hERG channels. Heteromeric channels consisting of WT and N-terminal truncated subunits have faster deactivation kinetics than WT hERG and are therefore more reminiscent of IKr (Pond et al., 2000). This alternatively spliced subunit called hERG1b, which possesses a unique stretch of 36 amino acids, is unable to form a functional homomeric channels (Lees-Miller et al., 1997; London et al., 1997; Robertson and January, 2006). Located at the C-terminus, the cyclic nucleotide binding domain (cNBD) is similar to that of pacemaker (HCN) channels in the heart.

Curiously, the cNBD has little effect on hERG channel gating as cAMP binding only induces a minor shift in the voltage dependence of activation (Cui, J. et al., 2000). Mutation or truncation of the C- terminus has far greater implications for channel processing in the ER and channel trafficking to the

PM (Akhavan et al., 2005).

1.3.2 hERG channel gating and kinetics

hERG channel kinetics are very unusual. Activation occurs with relatively slow kinetics, on the order of hundreds of ms to s, while inactivation kinetics are voltage dependent, occurring on the order of ms to tens of ms. As a result of these kinetics hERG is a delayed rectifier K+ channel with voltage-dependent activation (Sanguinetti et al., 1995). Little outward current is passed during depolarization because of rapid entry into the inactivate state. During repolarization, large outward tail currents are produced as

-26- the resting membrane potential is restored and hERG channels recover from inactivation before slowly deactivating.

The transmembrane electric field drives hERG channel activation by affecting the position of the positively charged S4 TM α-helices. hERG channel S4 TM domains possess a total of 7 positively charged Lys or Arg residues, located at roughly every 3rd position. Measurement of this voltage sensor movement reveals a small transient current which can be broken down into two kinetic components that differ by a factor of about 100. The slow kinetic component is associated with channel activation and specifically the movement of S4 domains, while the fast component is believed to be related to the movement of an inactivation voltage sensor (Vandenberg, J. et al., 2004). hERG channels must overcome a large energy barrier to open. This may be partially explained by the presence of numerous negative charges located on acidic residues on S1-S3. These residues form salt bridges with specific basic S4 residues, acting to stabilize the closed, intermediate and open states (Larsson et al.,

1996). Two particularly important Asp residues are located on the external side of the PM on S2 and

S3. These residues are sensitive to divalent cations which prevent the formation of salt bridges and shift the voltage-dependence of hERG activation to more positive potentials (Fernandez, D. et al.,

2005). Based on studies of Shaker channels, it is believed that movement of the hERG voltage sensor is electromechanically coupled to channel opening via the S4-S5 linker, the “activation gate”, which consists of an amphipathic α-helix located parallel to the PM (Tseng, 2001). This linker interacts with the C-terminal portion of the S6 on the same subunit, and stimulates channel opening and closing via a lever mechanism (Long et al., 2005a). Alternatively, the slow kinetics of hERG channel deactivation may be attributed to the PAS domain. Interaction of the N-terminal PAS domain with the S4-S5 linker stabilizes the hERG channel in the open state (Wang, J. et al., 1998). Deletion of the PAS domain results in a ten-fold acceleration of hERG channel deactivation kinetics.

hERG channels are also distinguished from other KV channels by their inwardly rectifying currents (of course hERG channels are not true inward rectifiers) (Smith et al., 1996). As hERG channels are

-27- depolarized to progressively more positive potentials, outward current is limited. hERG channel inactivation is voltage-sensitive occurring with rapid kinetics. Inactivation operates via a “C-type” mechanism, involving a slight constriction of the selectivity filter causing pore occlusion (Kiss and

Korn, 1998). This likely occurs when the outer most K+-binding site is unoccupied. C-type inactivation can be completely abolished by the mutation Ser631Val and mutations in the pore-loop and the N- terminal half of the S6 domain can also disrupt this fast inactivation process (Schonherr and

Heinemann, 1996; Fan et al., 1999). The rapid kinetics of hERG channel inactivation and recovery may be attributed to a more flexible and lengthy pore-loop and a narrower outer pore diameter, thereby requiring a smaller molecular motion.

1.3.3 Native hERG currents

The rapidly activating delayed rectifier current (IKr) can be distinguished from other repolarizing currents in the heart by its unique activation kinetics and its specific pharmacology (Sanguinetti and

Jurkiewicz, 1990; Sanguinetti, 1999). IKr currents activate more rapidly than IKs, but not faster than IKur.

IKr current can be specifically blocked by methanesulfonanilide anti-arrhythmic agents including E-

4031 (Mitcheson and Sanguinetti, 1999). Whole-cell patch clamp analysis of isolated guinea pig cardiomyocytes allowed for the first description of native IKr (Sanguinetti and Jurkiewicz, 1990).

Relative to IKs, IKr rapidly activates as demonstrated by a steep activation curve slope and a voltage required for half activation of current (V1/2) of -21.5 mV (vs. IKs V1/2 + 15.7 mV) (Sanguinetti and

Jurkiewicz, 1990). Time constants for IKr activation and deactivation as a function of membrane potential are bell-shaped, peaking between -30 and -40 mV at 170 ms. IKr inward rectification occurs at test potentials > -50 mV, resulting in a voltage-dependent decrease in peak current amplitude at potentials positive to 0 mV. This initial characterization demonstrated that IKr and IKs contribute equally to current amplitude during the plateau phase of the cardiac AP, as tested by 225 ms pulses with test potentials ranging from -20 to 20 mV (Sanguinetti and Jurkiewicz, 1990).

-28-

The hERG gene was identified during a high-stringency screen of the human hippocampus cDNA library, revealing a gene coding for a 1159 residue protein with a predicted molecular weight of 127 kDa (Warmke and Ganetzky, 1994). Mutations in the hERG gene were identified as causing LQT2 using the candidate gene approach in patients presenting with ventricular arrhythmia and sudden death

(Curran et al., 1995). hERG was subsequently cloned and demonstrated to code for the pore-forming

α-subunit of IKr in the heart (Sanguinetti et al., 1995; Trudeau et al., 1995). The initial characterization of hERG channel current was made possible by injecting Xenopus oocytes with hERG cRNA. In this overexpression system, whole-cell hERG currents have an activation V1/2 of -15 mV, obtaining peak outward conductances at roughly 0 mV (Sanguinetti et al., 1995). Native IKr/hERG currents have subsequently been recorded from isolated cardiomyocytes obtained from mouse, rat, pig, rabbit, dog and human (Carmeliet, 1992; Carmeliet, 1993; Wang, Z. et al., 1993; Wang, Z. et al., 1994; Liu and

Antzelevitch, 1995; Pond et al., 2000). Interestingly, the kinetics of IKr activation and deactivation are approximately 10-fold faster than hERG channels expressed in mammalian cells (Sanguinetti, 1999).

Identification of an N-terminal truncated hERG splice-variant highly expressed throughout the myocardium has shed some light on this discrepancy (Lees-Miller et al., 1997; London et al., 1997;

Jones et al., 2004). Although this splice variant does not produce functional hERG channels when expressed alone, it does form functional channels when heterologously expressed with WT hERG channels (Jones et al., 2004). The activation and deactivation kinetics of hERG/hERG1b channels almost identically recapitulate native IKr.

While it is widely accepted that hERG channels represent the pore-forming α-subunits of IKr in the heart, they are unable to fully reproduce that current when expressed heterologously. This led to an alternative hypothesis that like KvLQT1 and minK, hERG may interact with a β-subunit to form the channel complex underlying IKr. MinK-related peptide 1 (MiRP1) is a short 123 amino acid integral membrane protein which coassembles to form stable complexes with hERG, having the ability to modulate its expression and gating (Abbott et al., 1999). When heterologously coexpressed, MiRP1 reduces hERG channel trafficking to the PM, alters its pharmacological sensitivity, reduces single

-29- channel conductances and accelerates the rate of channel deactivation (Abbott et al., 1999). These results have not been confirmed in vivo, however, mutations in MiRP1 affecting hERG channel sensitivity to drug block hint at their potential relationship in human cardiomyocytes (Sesti et al.,

2000). In another study, heterologous expression of hERG, minK and MiRP1 demonstrated that hERG preferentially coimmunoprecipitated with minK which increased the rate of hERG channel trafficking to the PM, and that both proteins may be involved in the regulation of hERG channel trafficking rates

(Um and Mcdonald, 2007). The relationship between hERG channels and their putative β-subunits may be further complicated by the emergence of KCR1, a 12-TM domain subunit which has been demonstrated to alter the influence of MiRP1 on hERG drug sensitivity (Kupershmidt et al., 2003;

Nakajima et al., 2007). Finally, the heterogeneous expression patterns of hERG channels and their associated proteins throughout the heart allow for the fine tuning of hERG channel expression and modulation and make it even more complicated to make general conclusions regarding the nature of native hERG currents.

1.3.4 hERG channel pharmacology

hERG channels are remarkably sensitive to block by a wide variety of compounds, including psychiatric, antimicrobial and antihistamine drugs, as well some anti-arrhythmic agents (Sanguinetti and Tristani-Firouzi, 2006). Block of IKr/hERG channels by these drugs represent the predominant mechanism by which acquired LQTS operates. Dofetilide and quinidine are anti-arrhythmic agents used in the treatment of atrial arrhythmias but have the unwanted side effect of inducing ventricular arrhythmias and TdP in 2-7% of recipients (Sanguinetti et al., 1995; Camm et al., 2000). Cisapride, used for treating diseases of the GI tract, and terfenadine, an antihistamine, were both shown to cause LQTS in roughly 1/120,000 patients. In addition to these drugs, the use of numerous hERG-blocking compounds has been restricted or banned because of their dangerous side-effects (De Bruin et al.,

2005). Understanding why acquired LQTS can be induced in a small portion of the population has led to the hypothesis that normal physiology includes a built-in “repolarization reserve” characterized by

-30- redundancy of K+ channels and their normal level of expression (Roden and Spooner, 1999; Marbán,

2002). Acquired LQTS caused by hERG-channel blockers, or drugs which disrupt channel trafficking, or result in undesirable drug-channel interactions resulting in reduced hERG channel amplitude may act to reveal a sub-population possessing a reduced repolarization reserve. Therefore, certain polymorphisms may decrease channel expression or increase drug binding efficiency, resulting in an increased risk of LQTS (Roden, 2001).

Understanding the structural basis for hERG channel block has important implications for the design and evaluation of new experimental compounds. In fact, during preclinical assessment of most new compounds, hERG-screening is commonplace with the recent development of high-throughput planar patch-clamping (Dubin et al., 2005; Sanguinetti and Mitcheson, 2005). Alanine-scanning mutagenesis experiments have revealed specific residues that predispose hERG channels to drug block where other K+ channels are completely insensitive (Mitcheson et al., 2000). In particular, two highly conserved polar pore-helix residues, Thr623 and Ser624, and two unique S6-domain aromatic residues, Tyr652 and Phe656, are involved in drug binding (Lees-Miller et al., 2000; Mitcheson et al.,

2000; Laine et al., 2004). Mutation of these residues greatly reduces the affinity of the anti-arrhythmic compounds MK-499, cisapride and terfenadine (Sanguinetti and Mitcheson, 2005). Of particular importance for the hERG-drug interactions are the aromatic residues which are located on each hERG

α-subunit, pointing into the inner vestibule and producing a ring-structure for interaction (Fernandez,

D. et al., 2004). Interestingly, most of the compounds known to inhibit hERG channels possess one or more aromatic ring structures, which would promote a π-stacking interaction between the drug and

Tyr652 and Phe656 (Vandenberg, J. I. et al., 2001). hERG-channel block occurs from the intracellular side and only while the channel is in the open state (Snyders and Chaudhary, 1996; Spector et al.,

1996). Unbinding is a slow process, and is incomplete at negative potentials, implying that drug

“trapping” is common. This mechanism of drug-binding is enhanced by a larger inner vestibule, relative to other K+ channels, the result of an absent S6-domain Pro hinge (Mitcheson et al., 2000).

Drugs become trapped within the flexible vestibule without affecting deactivation kinetics.

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Understanding the exquisite drug-sensitivity of hERG channels has led to improvements in the development of new drugs, minimizing their off-target effects, as well as improving our general understanding of hERG channel structure and function.

1.3.5 Posttranslational processing of hERG channels

To date, 291 LQT2-linked mutations have been identified in hERG channels, the majority of these representing single amino acid substitutions yielding missense channel protein (see http://www.fsm.it/cardmoc/) (Anderson et al., 2006). Characterization of numerous such mutant hERG channels has overwhelmingly illustrated the consistent finding that LQT2 mutations cause a reduction in macroscopic hERG channel current. Interestingly, the mechanisms by which these mutations reduce or in many cases abolish hERG currents are numerous. hERG whole cell currents (IhERG) are the product of three factors: the number of functional channels inserted into the PM (N); the probability that an individual channel is open (PO); and the amplitude of single-channel conductances (i) (Delisle et al., 2004). Changes in the number of functional channels (N) at the PM can be the result of mutations causing irregular channel synthesis, for example, reduced transcription efficiency, translation errors, or other protein maturation abnormalities (class 1 mechanism). Additionally, reduction in the number of mature channels at the PM can be the result of impaired protein trafficking as in the case of LQT2 mutations affecting the hERG cNBD domain (class 2 mechanism) (Zhou et al., 1998a; Akhavan et al.,

2005). Changes in the open probability of a channel (PO) occur as a result of mutations affecting hERG channel gating or kinetics (class 3 mechanism). Modifying single-channel conductances (i) generally involves precise mutations in the pore region of the channel which affect permeability or selectivity of

K+ ions (class 4 mechanism). Finally, all three of these parameters can be affected when mutant hERG

α-subunits coassemble with WT subunits to form heteromeric channels (Fig. 8).

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Figure 8. LQT2 mutations cause a reduction in hERG channel whole cell current (IhERG)

Macroscopic IhERG is the product of the total number of channels at the PM (N), the probability that a channel is open (PO), and the single-channel conductance (i). hERG channels are transcribed in the nucleus, and mRNA is translated on ribosomes. In the ER hERG protein is core-glycosylated, folded and multiple subunits are assembled. Next, proteins are exported to the Golgi where complex glycosylation and sorting occurs. Vesicles containing hERG channels bud from the Golgi, traffic and insert into the PM. LQT2-causing mutations of hERG channels can disrupt any of these steps resulting in a reduction in (N), (PO), or (i), or combinations of these factors, thereby reducing IhERG.

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LQT2-linked hERG mutations are found throughout the channel structure, including both N- and C- termini, TM domains and the pore region. In order to characterize these types of mutations, a multi- faceted approach requiring the use of biochemical, molecular, immunocytochemical and electrophysiological techniques is required (Delisle et al., 2004). When studying these types of mutant hERG channels, a reduction in the PM expression of these mutants is the key to distinguishing them from WT hERG channels. Electrophysiological recordings generally show little or no detectable current, while Western blot analysis yields one band, corresponding to a core-glycosylated immature form of the hERG protein at 135 kDa. (Mature hERG protein undergoes complex glycosylation in the

Golgi apparatus yielding a 155 kDa band - a prerequisite for trafficking to the PM).

Immunocytochemistry reveals that many LQT2 mutants are not expressed at the PM, and are predominantly distributed in the perinuclear space. Overall, these results suggest that most trafficking-deficient hERG proteins are retained by the ER as a core-glycosylated protein.

hERG channel maturation through the secretory pathway is a complex process with many checkpoints ensuring quality control along the way to forming a mature protein. Following hERG channel synthesis and core glycosylation in the ER, immature hERG protein is transported to the Golgi apparatus where complex glycosylation occurs. Here, molecular chaperones such as heat conjugate/stress-activated protein 70 (Hc/sp 70) and heat shock protein 90 (Hsp 90) assist in proper hERG channel folding, or assist in the degradation of misfolded proteins. These two chaperones have been shown to coimmunoprecipitate with immature, core-glycosylated hERG protein. The role of

Hc/sp 70 is to assist in proper folding by binding to hydrophobic regions of the cytoplasmic side of the hERG channels during intermediate steps in protein folding. Hsp 90 is involved in preventing misfolded proteins from aggregating, and the inhibition of this chaperone has been shown to impair

WT hERG channel maturation, cell surface expression, and promote the poly-ubiquination pathway

(Ficker et al., 2003). Most recently, overexpressed chaperone protein FKB 38 (38-kDa FK506-binding protein, FKBP8) was shown to coimmunoprecipitate with immature hERG and rescue the trafficking- deficient hERG mutant F805C, while having no effect on WT hERG channels (Walker et al., 2007). hERG

-34- channels have multiple exit strategies from the ER and so modulating the balance of ER chaperone proteins may be the key to individualized treatment for some LQT2 mutations.

Forward transport through the secretory pathway is dependent on proper protein folding. hERG channels possess ER retention sequences which are masked following appropriate folding. If an ER retention sequence is detected during quality control, retrograde transport back to the ER may occur. hERG channels possess a putative ER retention sequence in the C-terminal at positions 1005-1007 with the sequence Arg-X-Arg (Kupershmidt et al., 2002). Truncation of the last 147 residues (downstream of the putative retention sequence) resulted in ER retention of the hERG protein. Additionally, deletion of residues 860-899 resulted in intracellular retention of the truncated protein (Akhavan et al., 2003).

These residues may play an important role in channel trafficking and folding, or they may be a part of a larger domain involved in channel maturation.

Once hERG channels have successfully folded and exited the ER, they enter the Golgi apparatus where they undergo complex glycosylation. hERG channels possess multiple consensus sites for N-linked glycosylation (Asn-X-Ser/Thr), however, residue N598 is the only residue that undergoes both core and complex glycosylation (Gong et al., 2002). hERG channel subunits are first synthesized as a 132 kDa peptide and then quickly undergo core-glycosylation in the ER to become a 135 kDa protein (Zhou et al., 1998b). These core-glycosylated immature hERG proteins are sensitive to digestion by endoglycosidase (Endo) H, producing the original 132 kDa peptide. Complex glycosylation yields a

155 kDa protein per subunit. This has been shown to be important for proper folding, export to the

PM, modifying protein function and improving protein stability (Petrecca et al., 1999; Delisle et al.,

2004). The time course of hERG channel maturation has been directly measured using pulse-chase metabolic labeling (Gong et al., 2002). Appearance of the 155 kDa, Endo H-resistant 155 kDa band demonstrates that hERG channels reach maturity in approximately 24 h at 37 °C (Robertson and

January, 2006).

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1.3.6 Rescue of mutant hERG channels

A large proportion of the 291 known LQT2-causing hERG mutations result in a channel trafficking- deficiency phenotype (Anderson et al., 2006). Recent advances in our understanding of how such mutations affect the quality control mechanisms and trafficking of hERG channels during normal protein maturation has led to the discovery that trafficking of many LQT2 mutants can be rescued under the right conditions in a mammalian cell system (Delisle et al., 2004). Work related to cystic fibrosis (CF), in which mutations of the CF transmembrane conductance regulator (CFTR) gene interferes with normal protein folding resulting in ER retention and rapid degradation, forms the basis for LQT2-rescue research. CFTR encodes a PM Cl- channel, with the most common CF mutation (70% of all cases) resulting from a single-point mutation ∆F508 (Cheng et al., 1990). Initial work focusing on restoration of normal protein trafficking revealed that ∆F508 trafficking could be rescued following incubation of mammalian cells over-expressing the mutant protein at a reduced temperature

(Denning et al., 1992). Subsequent work in this field led to the observation that trafficking of ∆F508

CFTR channels could be rescued following incubation with a variety of pharmacological agents which act as chaperones permitting misfolded proteins to acquire a folding configuration required for trafficking. These compounds include glycerol, trimethylamine N-oxide, sodium 4-phenylbutyrate, and deuterated water (Cheng et al., 1995; Brown et al., 1996; Sato et al., 1996). These findings have not only encouraged investigators to seek compounds used for the treatment of CF, but have also inspired researchers in the field of congenital LQTS.

Trafficking of numerous mutant hERG channels has been achieved following incubation at reduced temperature (Delisle et al., 2004). In one study, 16 of the 28 trafficking-deficient mutant channels had enhanced PM expression following incubation at reduced temperature (Anderson et al., 2006). Lower temperatures restore normal channel oligomerization, supporting export of mutant protein out of the

ER where it is normally retained. Additionally, reduced temperature promotes native folding by stabilizing intermediate steps in the protein-folding pathway (Delisle et al., 2004). While it has become

-36- generally recognized that a reduction in incubation temperature boosts the functional expression of difficult-to-express membrane proteins, this option is not clinically feasible. Of interest from an experimental point of view, expression of WT and mutant hERG channels at 30°C yields the greatest amount of surface-associated hERG, as measured by patch-clamp analysis, making these conditions favorable for high-throughput compound screening (Chen et al., 2007).

A variety of compounds including high-affinity hERG channel blockers, and hERG channel chaperone proteins have also been shown to restore trafficking in a variety of LQT2 mutants. Glycerol, incubated a molar concentrations, was shown to improve channel trafficking in the N470D hERG mutant (Zhou et al., 1999). It is likely that glycerol is able to stabilize an intermediate conformation of the immature hERG protein, allowing proper folding to occur (Sato et al., 1996). Alternatively, E-4031, cisapride, astemizole, quinidine, and fexofenadine are compounds which bind to and block hERG channels. All of these compounds have been shown to rescue hERG channel trafficking in numerous LQT2 mutations (Delisle et al., 2004; Gong et al., 2004). Perhaps the most thoroughly characterized compound is E-4031, a methanesulfonanilide drug which blocks IKr/hERG currents with high affinity

(Sanguinetti and Jurkiewicz, 1990). Micro-molar concentrations of E-4031 were introduced into the cell culture medium of mammalian cells over-expressing trafficking-deficient LQT2 mutant channels.

Out of the 28 mutants, trafficking was restored in 17 of those incubated with E-4031 (Anderson et al.,

2006). As mentioned previously, hERG channels are extremely sensitive to block by a variety of compounds which interact with aromatic residues located in the inner vestibule region of the channel.

Although the mechanism by which rescue occurs is not well understood, it is believed to involve drug binding to the inner vestibule of the hERG channel pore region, thereby stabilizing intermediate configurations of protein folding and promoting oligomeric stability (Ficker et al., 2002).

Although drug block is reversible, rescue of hERG channel trafficking by high-affinity blockers is not therapeutically feasible unless rescued IKr channels are sufficiently functional to support normal cardiac repolarization (Rajamani et al., 2002; Robertson and January, 2006). Fexofenadine is a

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derivative of terfenadine, a high-affinity hERG channel blocker, which has a 300-fold lower EC50 for hERG block (Rajamani et al., 2002). This compound has been shown to rescue trafficking of several hERG channels with LQT2 mutations, restoring normal IKr without affecting gating or permeation

(Zhou et al., 1999). Alternatively, thapsigargin has been shown to rescue numerous LQT2 mutations independently of the pore-blocking compounds (Delisle et al., 2003). Thapsigargin is a sarcoplasmic/endoplasmic reticulum Ca-ATPase (SERCA) inhibitor whose rescue mechanism is only speculative. It is believed that this drug acts to modulate luminal ER [Ca2+], affecting protein folding and Ca2+-dependent chaperone proteins (Delisle et al., 2004). Interestingly, thapsigargin has also been shown to restore cell surface expression of some CFTR mutants (Egan et al., 2002). Misfolded CFTR proteins are “handcuffed” by Ca2+-dependent chaperone proteins while they await degradation. A

SERCA-mediated reduction in intracellular Ca2+ causes these proteins to disassociate from the misfolded proteins, thereby allowing their escape to the PM (Robertson and January, 2006).

Generally, rescue of channel trafficking interrupts degradation of incorrectly folded proteins which have been sorted by cellular quality control mechanisms. Compounds including chemical chaperones and pore blockers act to increase the likelihood of reaching a native protein conformation, and help to stabilize intermediate protein configurations. This serves to promote proper channel folding and oligomerization, however, each LQT2 mutant affects posttranslational processing of hERG channels differently, and so incubation with a particular compound will not restore trafficking in every mutation

(Anderson et al., 2006). Finally, the aforementioned compounds do not affect WT hERG channel trafficking or PM expression, a testament to the highly-tuned quality control mechanisms inherent in the production of normal hERG channels (Delisle et al., 2004).

1.4 SNARE proteins

SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) proteins are most widely recognized as components of the protein complexes which underlie membrane fusion events.

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Cellular communication between distinct membrane-enclosed organelles in eukaryotic cells occurs via the exchange of trafficking vesicles between these distinct compartments. SNAREs represent a highly conserved family of proteins, present on both the vesicular and target compartment membranes, which drive membrane fusion by utilizing the free energy released during formation of a four-helix bundle. These basic cellular events underlie a wide variety of essential biological mechanisms including cell growth, membrane repair, cytokinesis, exocytosis and synaptic transmission (Jahn and Scheller, 2006). More recently, it has become apparent that outside of their well characterized complex-forming role in membrane fusion, many SNARE proteins also functionally interact with and modulate the expression and function of a variety of ion channel types (Peters et al.,

2001; Jarvis and Zamponi, 2005; Leung et al., 2007). These include K+ ion channels expressed in the heart (Kang et al., 2004; Kang et al., 2006; Yamakawa et al., 2007; Ng et al., 2008).

1.4.1 Structure of SNARE proteins and mechanism of membrane fusion

SNARE protein expression is highly abundant, representing 1% of total brain protein, more than the expression of all ion channels combined (Walch-Solimena et al., 1995). Characterized by a simple domain structure and each containing a common 60-70 amino acid “SNARE motif”, there are 36 known membranes of this protein superfamily in humans (Jahn and Scheller, 2006). Originally, SNARE proteins were classified into two groups: the v-SNAREs (vesicle-membrane SNAREs) and the t-SNARES

(target-membrane SNAREs). This classification scheme reflected the belief that distinct SNARE proteins participate in fusion reactions in specific membrane orientations. In fact, many SNARE proteins are capable of forming functional SNARE complexes from both “donor” and “acceptor” membranes and can also function in both anterograde and retrograde directions (Sollner et al., 1993).

The current classification scheme for SNARE proteins reflects the fact that SNARE complexes are composed of 4 intertwined parallel α-helices, each of which is provided by a different SNARE motif.

Individual monomeric SNARE proteins do not have structured SNARE motifs, however, when combined they spontaneously form extremely stable helical core complexes (Fasshauer, 2003). The

-39- center of this complex consists of 16 stacked layers of hydrophobic interacting side chains, except

“layer 0” which contains 3 highly conserved Glu and 1 Arg residues. These structural features gave rise to the current classification system for SNARE proteins – Qa-, Qb-, Qc- (Fasshauer et al., 1998). Each component is required for membrane fusion, and is highly conserved throughout evolution (Bock et al., 2001).

Most SNARE proteins possess one TM domain located at the C-terminal. A short rigid linker joins the

TM domain to the SNARE motif, causing it to stand upright relative to the membrane (Kiessling and

Tamm, 2003). This ensures that during SNARE core complex formation, “zippering” of the 4 α-helices proceeds from C-terminus towards the N-terminus, thereby transferring energy caused by the straining of the SNARE motifs onto membranes, bending them or disturbing the hydrophilic/hydrophobic boundary (Jahn and Scheller, 2006). This is believed to press the opposing membranes together, deforming them and facilitating the formation of fusion stalks which precede hemifusion and fusion-pore opening. The number of SNARE complexes required for a single fusion event to occur is unclear, however, estimates range between 3 and 15 (Montecucco et al., 2005).

Disassembly of SNARE core complexes is facilitated by NSF (N-ethylmaleimide-sensitive factor) which transforms the complex from a trans- to a cis-configuration. These complexes normally remain stable under conditions as extreme as 80 °C and 2M SDS (Fasshauer et al., 2002).

X-ray crystallographic analysis has revealed the exquisite detail of monomeric SNARE proteins and heteromeric SNARE complexes (Fernandez, I. et al., 1998; Sutton et al., 1998; Misura et al., 2000; Antonin et al., 2002). Fig. 9 illustrates the characteristic domain structure of SNARE protein subfamilies, as well as the crystal structure of the neuronal SNARE core complex which contains the SNARE motifs of three proteins: the Qa-SNARE syntaxin 1A (STX1A), the Qbc-SNARE SNAP-25 (synaptosome-associated protein of 25 kDa), and the R-SNARE VAMP (vesicle-associated membrane protein)/synaptobrevin.

SNAP-25, is classified as a Qbc-SNARE because it does not possess a TM domain, and interestingly, it possesses 2 SNARE motifs which are required for neuronal core complex formation. The Qa-SNARE

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Figure 9. Structure and assembly of SNARE proteins

A) Schematic diagram illustrating the domain structure of the SNARE protein subfamilies. All subfamilies contain a SNARE motif which is involved in forming the SNARE core complex during membrane fusion. Qa-SNAREs include STX1A and have an N-terminal 3-helix bundle which is loosely linked to the SNARE motif. Qb-, Qc- and R- SNARES may have a variety of types of N-terminal domains (oval shapes). The Qbc-SNARE subfamily is comprised of SNAP-25 which possesses 2 SNARE motifs and no TM domain. Dashed lines around domains indicate that they are not expressed in every member of the subfamily. B) Ribbon diagram obtained from the three dimensional crystal structure of the neuronal SNARE core complex. The core complex is comprised of the proteins STX1A, SNAP-25 and VAMP which contribute 4 α-helical SNARE motifs to the complex. C) Ribbon diagrams obtained from the three dimensional crystal structure of the STX1A N-terminal. The HABC domain (left) is also shown interacting with the H3 domain (SNARE motif, right) which produces the STX1A closed conformation. This structure was solved as a component of the Munc18-syntaxin-1 complex (Misura et al., 2000).

Figure adapted with permission from Macmillan Publishers Ltd: Nature Reviews Molecular Cell Biology (Jahn and Scheller, 2006), copyright (2006).

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STX1A has a large N-terminal domain, connected to the SNARE motif by a long and flexible linker

(Fernandez, I. et al., 1998). Known as the HABC domain, a complex of 3 anti-parallel α-helical bundles can reversibly associate with the SNARE motif (called the H3 domain in STX1A), forming a “closed” conformation and yielding the protein inactivate (Misura et al., 2000). Uninhibited STX1A proteins are believed to adopt a dimer consisting of 2 identical proteins with interacting SNARE motifs (Lerman et al., 2000). Formation of homomeric SNARE complexes may be prevented by the disassembly protein

NSF. N-terminal domains in SNARE proteins are highly variable, and are not necessarily required for normal protein function (Jahn and Scheller, 2006). These domains may also function as platforms for the binding of additional proteins required for membrane fusion as in the case of Sec1/Munc18- related (SM) proteins, or for the direct regulation of various SNAREs (Toonen and Verhage, 2003).

SNARE proteins, although small and composed of simple domain-structures, are the vivacious molecular machines required for intracellular fusion events. While intensive research continues to clarify the complex reaction cycle of membrane fusion, it is becoming increasingly clear that uncomplexed monomeric SNAREs may have numerous alternative cellular functions.

1.4.2 Modulation of ion channels by SNARE proteins

A wide variety of diverse proteins bind to SNARE proteins, in fact, over 100 binding partners have been reported for synaptic SNARES alone (Jahn & Scheller, 2006). Many of these proteins regulate the expression and function of monomeric “free” SNAREs, affecting sorting and recycling, recruitment into trafficking vesicles, the formation of docking complexes, and the regulation of SNARE motif capability

(Peden et al., 2001; Siniossoglou and Pelham, 2001; Hu et al., 2002; Pennuto et al., 2003; Collins et al.,

2005). Alternatively, SNARE proteins bind to voltage-gated ion channels in neuronal and neuroendocrine cell types, functionally coupling the proteins and altering channel gating and expression.

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STX1A and SNAP-25 both bind to a common motif on Ca2+ channels called the synaptic protein interaction site (synprint) motif which is located on the II-III linker region of α1-subunits (Leveque et al.,

1994; Sheng et al., 1994; Wiser et al., 1999). Both STX1A and SNAP-25 bind to N-type (CaV2.2) and P/Q- type (CaV2.1) CaV channels with high affinity causing changes in channel gating and expression

(Bezprozvanny et al., 1995; Rettig et al., 1996). STX1A specifically reduces N-type CaV current amplitude and stabilizes channel inactivation (Bezprozvanny et al., 1995; Wiser et al., 1999). Additionally, STX1A and SNAP-25 independently bind with lower affinities to the synprint motifs on the L-type (CaV1.2 and

CaV1.3) CaV channels altering channel gating (Wiser et al., 1996; Yang et al., 1999). Binding of STX1A to

2+ 2+ the L-type Ca channel (CaV1.2) causes local increases in Ca current which affects synaptic vesicle fusion with the target membrane (Cohen et al., 2003; Kobayashi et al., 2007). STX1A is also capable of cross-talk with G-proteins and may enhance G-protein inhibition of Ca2+ channels (Stanley and

Mirotznik, 1997; Jarvis et al., 2000; Jarvis and Zamponi, 2001a). Thus SNARE proteins not only serve to regulate membrane fusion and exocytosis in neuroendocrine cells, but also serve as sites for feedback inhibition (Jarvis and Zamponi, 2005). Coexpression of STX1A and SNAP-25 yielding a stable binary

SNARE complex reverses the inhibitory effects of these proteins on L-, N-, and P/Q-type CaV channels

(Tobi et al., 1998; Zhong et al., 1999). SNARE protein-mediated regulation of Ca2+ channels represents a direct mechanism which influences a variety of Ca2+ channel types, and is significant because of the tremendous potential for the regulation of synaptic Ca2+ and the tuning of neurotransmission, as well as the regulation of exocytosis (Jarvis and Zamponi, 2001b; Zamponi, 2003).

STX1A and SNAP-25 have also been shown to bind to CFTR Cl- channels as well as amiloride-sensitive epithelial Na+ channels (ENaC), affecting channel trafficking and gating (Naren et al., 1997; Saxena et al., 1999; Vankeerberghen et al., 2002). STX1A inhibits both CFTR and ENaC current amplitudes, while a homologous SNARE protein Syntaxin 3 (STX3) increases the current amplitude of these channels

(Naren et al., 1997; Saxena et al., 1999). STX1A directly binds to a regulatory domain consisting of acidic residues located on the N-terminal of CFTR. STX1A inhibits cAMP-dependent trafficking of the channel to the PM, and also prevents cAMP-dependent activation of Cl- transport (Naren et al., 1997).

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Phosphorylation of the regulatory domain by PKA abolishes this interaction. SNAP-25 also interacts with the CFTR regulatory domain, and works cooperatively with STX1A to inhibit channel function

(Naren et al., 2000). These results illustrate the ability of SNARE proteins to have differential affects on multiple ion channels, as well as modulating their function differentially by interaction with multiple

SNARE protein isoforms.

The interaction of SNARE proteins and K+ channels represents a relatively new paradigm in the regulation of ion channel gating and expression in both secretory and non-secretory cells. In pancreatic islet β-cells, the SNARE protein STX1A has been shown to functionally interact with KV2.1 and the SUR2A subunit of the β-cell KATP channel to modify their gating behavior, thereby regulating cell excitability and insulin secretion (Leung et al., 2003; Cui, N. et al., 2004; Leung et al., 2007). STX1A strongly binds to the C-terminus of KV2.1, functionally inhibiting current amplitude by increasing the voltage sensitivity of steady-state inactivation and by slowing the kinetics of channel activation

(Leung et al., 2003). In contrast, STX1A binds to the N-terminus of KV1.1 and inhibits current amplitude by enhancing channel inactivation (Michaelevski et al., 2002; Michaelevski et al., 2003). Interestingly,

STX1A reduces the cell surface expression of both of these channels. The binding of SNAP-25 to KV1.1 and KV2.1 is in contrast to STX1A as this binding occurs at the N-terminus of both channels (Fili et al.,

2001; Ji et al., 2002b; He et al., 2008). SNAP-25 also binds to the KV1.1 β-subunit, however, this interaction occurs via the C-terminus (Michaelevski et al., 2002). Finally, STX1A was also found to inhibit KV1.2 current amplitude, slowing the kinetics of channel activation and right-shifting the activation curves (Neshatian et al., 2007). Paradoxically, STX1A actually increased KV1.2 channel trafficking and PM expression. The binding of STX1A and SNAP-25 to KV channel functional domains not only greatly affects channel gating and expression, but their binding to specific and distinct sites within and between the channels allows for a much more complicated and intricate control mechanism. This is in extreme contrast to the SNARE-mediated control of CaV channels which possess a common synprint motif and function relatively uniformly among the various CaV channel families.

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1.4.3 Interaction of SNARE proteins and cardiac potassium channels

The Tsushima and Gaisano laboratories have identified the expression of the SNARE proteins, STX1A and SNAP-25 in rat and mouse ventricular myocytes, and more importantly, the functional interaction of these SNARE proteins with cardiac KV4.2 and KATP channels (Kang et al., 2004; Kang et al., 2006;

Yamakawa et al., 2007; Ng et al., 2008). More recently, the expression of SNARE proteins in mouse heart was confirmed in both neonatal and adult atrial myocytes (Peters et al., 2006). Interestingly, the differential expression of neonatal and adult SNARE proteins appears to be isoform specific, with

STX1A and SNAP-25 expression not becoming predominant until adulthood. The functional role of

SNARE proteins in non-secretory excitable cells has not been fully elucidated. STX1A inhibits cardiac

KATP channels by acting on their SUR2A β-subunits (Kang et al., 2004). STX1A binding strength increases with progressively reduced pH, thereby increasing KATP channel inhibition (Kang et al., 2006).

This may serve as a protective mechanism during mild exercise or extreme cardiac ischemia such that inhibition of KATP currents would negate the induction of fatal reentrant arrhythmias. STX1A interacts preferentially with the N-terminus of KV4.2 causing a reduction in current amplitude, producing a depolarizing shift in the steady-state inactivation curve, and accelerating the rate of recovery from inactivation (Yamakawa et al., 2007). STX1A also markedly impaired KV4.2 channel trafficking to the plasma membrane, similar to the impairment of KV2.1 channel trafficking mentioned above. Recently

STX1A has also been shown to interact with cardiac KV4.3 channels (Ahmed et al., 2007). Surprisingly,

STX1A increased channel trafficking to the PM but inhibited KV4.3 current density by producing a slight hyperpolarizing shift in steady-state inactivation. Although this finding is in contrast to the affect of SNARE proteins on K+ channels, it serves to highlight the exquisite level control of channel expression and gating. While cardiac myocytes display abundant plasma membrane expression of

STX1A and SNAP-25, it is less clear whether they modulate ion channels as abundantly as in neuronal and neuroendocrine cells. Overall, these results suggest that SNARE proteins may underlie a novel endogenous control mechanism involved in the regulation of cardiac ion channel trafficking and gating.

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Chapter 2: Research Objectives

2.1 Rationale

The endogenous expression of the SNARE protein STX1 has been identified in rat ventricular cardiomyocytes by confocal microscopy and by the appearance of doublet bands using Western blot analysis of whole heart lysates, indicating the expression of STX1A and STX1B (Kang et al., 2004). This result has recently been recapitulated in mouse ventricular myocytes, in which the expression of

STX1A and STX1B were shown to be primarily plasma membrane-located (Ng et al., 2008). Another study identified the expression of numerous SNARE proteins in neonatal and adult atrial myocytes

(Peters et al., 2006). This study demonstrated an isoform shift in the expression of SNARE proteins from SNAP-23 and STX4 in neonatal myocytes to SNAP-25 and STX1A in adults. This suggests that the differential expression of SNARE proteins and their regulation of cardiac function may be age-related.

It has been well established that STX1A and other SNARE proteins are functionally coupled to the regulation of ion channel expression and gating in neuronal and neuroendocrine cell types (Wiser et al., 1999; Yang et al., 1999; Jarvis and Zamponi, 2005; Leung et al., 2007; Singer-Lahat et al., 2008).

+ STX1A has been shown to functionally interact with several K ion channels including KV1.1, KV1.2,

KV2.1 and KATP channels through the use of overexpression assays in mammalian cells or through the use of primary secretory cell types (Michaelevski et al., 2002; Leung et al., 2003; Cui, N. et al., 2004;

Neshatian et al., 2007). STX1A inhibited channel function in all of these K+ channels, although this was achieved by a variety of mechanisms and through binding to distinct regions of these channels. For example, STX1A reduced the cell surface expression of KV1.1 and KV2.1 (Michaelevski et al., 2002; Leung et al., 2005), but actually increased channel trafficking and PM expression in KV1.2 (Neshatian et al.,

2007). Additionally, STX1A enhanced KV1.1 and KV2.1 channel inactivation while slowing the activation kinetics of KV1.2 and KV2.1 (Leung et al., 2003; Michaelevski et al., 2003; Neshatian et al., 2007). Binding of STX1A occurred at the KV1.1 N-terminus, while binding of KV2.1 occurred at the C-terminus

-46-

(Michaelevski et al., 2002; Leung et al., 2003). The fact the STX1A can specifically and differentially regulate these K+ channels illustrates the fact that a common binding site does not exist, and so binding to different structural regions of the channels produce unique interactions which underlie specific alterations in channel expression and function.

STX1A has also been demonstrated to inhibit the function of cardiac K+ ion channels. STX1A inhibits the function of cardiac SUR2/KATP channels via interaction with the SUR2 β-subunit, and blocks acid pH-induced activation of these channels (Kang et al., 2004; Kang et al., 2006). Additionally, STX1A has been shown to reduce current density of cardiac KV4.2 and KV4.3 in Xenopus oocytes and HEK 293 cells, which are the main constituents of the transient outward K+ current in the heart (Ahmed et al., 2007;

Yamakawa et al., 2007). STX1A was found to bind to the KV4.2 N-terminus, reducing current amplitude by inhibiting channel trafficking. This was despite a STX1A-mediated right-shift in steady-state inactivation and enhancement of channel recovery from inactivation (Yamakawa et al., 2007).

Paradoxically, STX1A reduced KV4.3 current density despite increasing channel trafficking to the PM without affecting channel synthesis (Ahmed et al., 2007).

Based on these observations, we believe that SNARE proteins have fundamental cellular functions beyond membrane fusion events, involving the regulation of ion channel trafficking and gating in the heart. Furthermore, we believe that the SNARE protein STX1A may be involved in the regulation of

+ hERG channels which represent the rapidly-activating delayed rectifier K current (IKr) involved in phase 3 repolarization of the cardiac action potential. I will investigate the functional interaction of these proteins by utilizing a multi-disciplinary approach involving electrophysiological, molecular, and biochemical techniques. Therefore, the focus of this study is to determine the extent to which the

SNARE protein STX1A modulates the functionality of hERG channels.

-47-

2.2 Hypothesis

SNARE proteins in the heart carry out fundamental cellular duties, such as regulation of ion channel expression and function. Specifically, STX1A is an important intrinsic regulator of hERG channel trafficking and gating. These studies may have important implications in understanding the regulation of hERG channel trafficking and gating, which has clinical implications for heart disease and long QT-related syndromes in the heart.

2.3 Specific aims and experimental design

Aim 1 – To perform electrophysiological assessment of the hERG-STX1A interaction

The extent to which coexpression of STX1A modulates hERG current amplitude and gating will be assessed using whole-cell patch clamping. hERG channels will be heterologously expressed in the

HEK 293 mammalian cell line without or with STX1A. An exhaustive assessment of hERG channel gating and kinetics will be performed by utilizing specific voltage protocols designed to investigate these properties (Sanguinetti et al., 1995).

Aim 2 – To evaluate STX1A-induced changes in hERG trafficking, surface expression and localization

Changes in hERG channel trafficking and localization following cotransfection with STX1A will be qualitatively investigated using immunocytochemistry and confocal immunofluorescence. hERG channel maturation following cotransfection with STX1A will be assessed using Western blot analysis which reveals the expression of two protein bands, the larger band corresponding to mature, complex-glycosylated hERG protein (Zhou et al., 1998b).

Aim 3 – To determine the site of hERG-STX1A binding

Assessment of hERG and STX1A binding in our overexpression system will be analyzed using GST pulldown and coimmunoprecipitation assays. Furthermore, I will utilize truncated GST-STX1A-fusion

-48- proteins in order to determine which STX1A domain is involved in hERG interaction (Cui, N. et al.,

2004). Additionally, I will utilize a variety of N- and C-terminal hERG truncation mutations in order to assess the site of STX1A interaction with hERG channels (Wang, J. et al., 1998; Akhavan et al., 2003).

This may reveal structural information regarding the potential protein-protein interaction and may correlate with functional data obtained from electrophysiological assessment.

Aim 4 – To disrupt STX1A-mediated changes in hERG channel function or expression

STX1A-mediated changes in hERG channel trafficking or gating could be caused by direct interaction of those proteins. Therefore I will assess whether modification of hERG channel function can be abolished through the use of hERG channel blockers or reduced temperature. Similar to rescue of

LQT2-hERG mutations, these methods may compete with or disrupt the effects of STX1A coexpression, therefore restoring normal hERG channel function (Delisle et al., 2004).

Aim 5 – To evaluate the endogenous expression of hERG and STX1A in cardiac myocytes and HL-1 cells

The physiological relevance of the potential hERG-STX1A interaction will be tested by determining whether these proteins are expressed in native rat and mouse myocytes, and in HL-1 cells, a mouse atrial myocyte tumor-derived cell line (Claycomb et al., 1998). Additionally, the suitability for these model systems for further experimentation will be assessed.

2.4 Relevance

The results of this study will serve to not only provide valuable insight into the role of SNARE proteins in the heart, but there are also implications for LQT-related syndromes. hERG channel dysfunction impairs repolarization during phase 3 of the cardiac action potential, causing prolonged AP duration leading to ventricular fibrillation and deadly torsade des pointes (Curran et al., 1995). Currently, 30-35% of patients diagnosed with LQTS cannot be linked to any of the known genes (Crotti et al., 2008).

STX1A-impairment of hERG channels could represent a new paradigm of control in healthy and

-49- pathological human cardiac physiology. More generally, this study may contribute to a better understanding of an emerging biological mechanism involving the exquisitely fine-tuned SNARE protein-mediated regulation of KV channels throughout the body, and particularly in the heart.

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Chapter 3: Materials and Methods

3.1 DNA Constructs

hERG pSP64 DNA was generously provided by Dr. Michael Sanguinetti (University of Utah) and pCMV-

STX1A (WT) was from Richard Scheller (Genentech, San Francisco, CA) (Bennett et al., 1993; Sanguinetti et al., 1995). SNAP-25 was provided by the late Dr. Heiner Niemann (Medizinische Hoshschule,

Hanover, Germany) (Binz et al., 1994). The HA-tagged wild-type hERG construct as well as HA-tagged

C-terminal hERG truncation mutations ∆1120, ∆1045, ∆1000, ∆960, ∆899, ∆860, ∆814 and ∆860-899 were kindly provided by Dr. Alvin Shrier (McGill University, Montreal, QC) (Akhavan et al., 2003). The

HA-tagged WT construct was prepared using a vector that allowed for the addition of an HA- tag to the amino terminus of the target protein. hERG truncation mutations were named according to the last amino acid residue upstream of the truncated mutations. Deletions were numbered according to the specific residues that were removed. The N-terminal hERG truncation mutations ∆2-16 and ∆2-354 were provided by Dr. Gail Robertson (University of Wisconsin-Madison) (Wang, J. et al., 1998). We further modified these constructs through the addition of an N-terminal HA-tag. Channel constructs were confirmed by restriction enzyme analysis followed by sequencing and Western blot analysis. All channel constructs were subcloned into pcDNA3 (Invitrogen, Burlington, ON).

3.2 Generation of GST-fusion proteins

STX1A in pGEX-4T-1 was provided by Dr. William Trimble (Hospital for Sick Children, Toronto, ON).

GST fusion proteins were generated using pCMV-STX1A as a template for the production of full length

STX1A, STX1A-HABC (corresponding to amino acids 1-160) and STX1A-H3 (amino acids 191-256) which were then subcloned into a pGEX-4T1 vector (GE Healthcare, Baie d'Urfe, QC). All constructs were verified using DNA sequencing. GST fusion protein expression and purification were performed following the manufacturer’s instructions. STX1A was obtained by cleavage of GST-STX1A with

-51- thrombin (Sigma-Aldrich Canada, Oakville, ON) prior to elution of the GST fusion protein from glutathione-agarose beads.

3.3 Cell culture

tsA-201 cells were used for preliminary experiments and were grown at 37°C in a humidified 5% CO2 and 95% O2 incubator. Cell culture medium was Dulbecco’s Minimum Essential Medium (DMEM)

(Invitrogen) containing 4.5 g/L glucose and L-glutamine. TsA-201cells are HEK 293 cells stably transfected with a T-antigen which promotes the replication of certain viral promoters including CMV- containing constructs. We switched to the use of naïve HEK 293 cells as we found no drastic difference in transfection efficiency following electrophysiological assessment of ion channel function.

HEK 293 cells were grown under the same conditions as tsA-201 cells. Both cells lines were maintained in DMEM supplemented with 10% fetal bovine serum (FBS; Invitrogen) and penicillin/streptomycin (100 units/mL; 100 μg/mL; Invitrogen). Additionally, stably transfected hERG-

HEK 293 cells were obtained from Dr. Craig January (University of Wisconsin-Madison) and used for electrophysiological experiments (Zhou et al., 1998b). These cells were maintained identically to the cell types above, except for the exclusion of penicillin/streptomycin and the addition of 400 μg/mL geneticin (G418, Sigma) an aminoglycoside antibiotic which blocks polypeptide synthesis. hERG-HEK

293 cells possess a neomycin-resistant gene driven by the SV40 promoter from Tn5 encoding an aminoglycoside 3’-phosphotransferase, APH 3’ II which ensures that only transfected cells survive. In preparing the stably transfected cell line, Dr. January’s group picked single colonies of hERG-HEK 293 cells and assessed them in order to ensure the expression of robust hERG currents with minimal background currents which are predominant in tsA-201 and naïve HEK 293 cells. Cells were not used beyond 30 passages (3:1 split), and were never allowed to reach full confluency during culture.

HL-1 cells were obtained from Dr. William C. Claycomb (Louisiana State University, New Orleans, LA)

(Claycomb et al., 1998). These cells are a cardiac muscle cell line derived from the AT-1 mouse atrial

-52- cardiomyocyte tumor lineage. In addition to retaining their differentiated cardiac morphological, biochemical and electrophysiological characteristics, they are able to contract in culture. HL-1 cells were grown at 37°C in a 5% CO2 humidified incubator and fed Claycomb’s medium (SAFC Biosciences,

Lenexa, KS) supplemented with 10% FBS (Sigma), penicillin/streptomycin (100 μg/mL; Invitrogen), 0.1 mM norepinephrine (Sigma) and 2 mM L-glutamine (Sigma). Cells were passaged once they reached confluency and split 3:1 as lower cell densities may cause cells to lose their differentiated characteristics. Cells were grown on 100 mm dishes coated with 12.5 μg/mL fibronectin and 0.02% gelatin.

3.4 Transfection and drug treatment

Several different amounts of cDNA were tested to achieve optimal hERG expression as assessed by electrophysiological and molecular biology. For all experiments, 1.0 μg of hERG cDNA were used per

35 mm dish. A ratio of 2:1 (hERG: STX1A) cDNA was used such that 0.5 μg of STX1A cDNA were transfected per 35 mm dish. This ensured an excess amount of STX1A protein to hERG channels. hERG transfection yields an immature protein product of 135 kDa and a mature protein product of 155 kDa. Mature protein accounts for roughly 30% of the total hERG protein in our transfection system and each protein represents one subunit of a tetrameric channel, the approximate molecular weight of an average hERG channel is 564 kDa. The molecular weight of STX1A is 35 kDa. The following calculation illustrates how our system produces a roughly 8:1 molar excess of STX1A:

135 kDa x 4 subunits = 540 kDa x 70% = 378 kDa

155 kDa x 4 subunits = 620 kDa x 30% = 186 kDa

Average hERG channel = 378 kDa + 186 kDa = 564 kDa

STX1A = 35 kDa

Transfection ratio of 2 hERG : 1 STX

-53-

Therefore, ; x ≈ 8

1 hERG : 8 STX1A molar ratio

TsA-201 and HEK 293 cells used for electrophysiological experiments were grown in 35 mm dishes and were transiently transfected at greater than 50% confluency with green fluorescent protein (GFP) (0.3

μg) and hERG (1.0 μg) with or without STX1A (0.5 μg) using LipofectamineTM 2000 (Invitrogen) according to the manufacturer’s instructions. Twenty-four h after transfection, cells were trypsinized and placed in 35 mm dishes in low density and cultured overnight for single-cell electrophysiological recordings. Successful transfection was identified by visualizing a moderate amount of green fluorescence. hERG-HEK 293 cells were prepared in a similar fashion with or without transient GFP transfection in order to determine that GFP did not substantially alter hERG channel current properties. For electrophysiological experiments hERG-HEK 293 cells were transfected without or with

STX1A (0.5 μg) and GFP (0.3 μg). In another set of experiments, SNAP-25 (0.5 μg) transfection was substituted for STX1A (see Appendix 2). For biochemical experiments, cells were grown in 100 mm tissue culture dishes and the amount of DNA used was 6 times that mentioned above for each DNA species. Groups were transfected and harvested at similar confluences to ensure similar protein yield.

For immunofluorescence experiments, cells were grown on 18 mm glass coverslips.

In some experiments, cells were grown in the presence of 5 μM E-4031 (Calbiochem, San Diego, CA) for 24 h prior to harvesting for biochemical experiments or electrophysiological assessment.

Following E-4031 incubation, cells were washed with PBS and replaced with fresh E-4031-free medium four times over a 1 h period prior to being used for electrophysiological experiments. All electrophysiological recordings were performed within 1-2 h following removal from culture conditions. Tunicamycin (Sigma, T7765) was used to block the synthesis of N-linked glycoproteins.

Tunicamycin was diluted in distilled H2O to make a 10 mg/ml stock solution and was added to

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Dulbecco's modified Eagle's medium for 24-48 h at a concentration of 10 μg/ml prior to molecular biology experiments.

3.5 Electrophysiology

Voltage-gated K+ channel (hERG) recordings of single cells were performed using the whole- cell configuration of the patch clamp technique (Fig. 10). Recordings were made using a HEKA EPC-10 amplifier and Pulse software (HEKA Electronics Inc, Mahone Bay, NS). Pipettes were pulled from 1.5 mm borosilicate glass capillary tubes (World Precision Instruments, Sarasota, FL) using a programmable micropipette puller (Sutter Instrument, Novato, CA). Pipettes were heat polished and resistances were obtained ranging from 2-4 MΩ when filled with internal solution and measured using the EPC-10 and Pulse software. Internal (pipette) and external (bath) solutions were prepared and stored for use within 2 months. Components of these solutions are listed in Tables III and IV. Once whole-cell configuration was established (following achievement of a gigaohm (GΩ) seal), the cell was held at -80 mV and subjected to various experimental protocols as detailed in the Results section and legends. All recordings were performed at room temperature (~22 °C).

3.6 Western blot analysis

Cells were grown for 48 h on 100 mm plates to similar confluences. All steps were performed at 4 °C.

Cells were washed three times with ice cold PBS and incubated with 400 μL lysis buffer for 20 min.

Lysis buffer contained 0.5% Nonidet P-40 buffer; 50 mM Tris HCl (pH 8.0); 75 mM NaCl and a protease inhibitor cocktail (Roche Applied Science, Indianapolis, IN). Cell lysates were scraped from the dishes into 1.5 mL Eppendorff tubes, placed on ice for an additional 30 min with frequent vortexing. During this incubation time, cell lysates were passed through a 20G 1 ½” syringe 10 times in order to break up cells. Lysates were then centrifuged at 16,000 × g for 30 min and the DNA-containing pellet was discarded. Protein concentration was obtained using a modified Bradford method (BioRad assay kit,

-55-

Figure 10. Whole-cell mode of the patch clamp electrophysiology technique

Electrophysiological assessment of hERG channels was achieved using the whole-cell mode of the patch clamp technique. A thinly-pulled glass micropipette was filled with internal (pipette) solution containing 140 mM KCl and is installed on an electrode connected to a HEKA EPC-10 amplifier. HEK 293 cells can be transiently transfected with GFP so that they appear green under fluorescent light indicating a successfully transfection (i.e. in the case of STX1A cotransfection). Cells are grown in 35 mm dishes and culture medium is washed and replaced with an extracellular (bath) solution containing 4 mM KCl and 140 mM NaCl prior to use for patch clamp experiments. Whole-cell patch clamping involves “breaking in” to gain electrical access to the cell interior using negative pressure and obtaining a “Gigaohm seal”. This allows the experimenter to send voltage commands via computer software and the amplifier, through the electrode which fixes voltage and measures changes in current amplitude, sending these recordings back through the amplifier to be displayed and analyzed with computer software.

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Table III: Preparation of external solution for patch-clamp electrophysiology experiments

External (bath) solution components

Final Volume = 1 L (in ddH20)

Final Conc. Compound M.W. Weight/Vol

140 mM NaCl 58.44 g/M 8.1814 g 4 mM KCl 74.55 g/M 0.2982 g

1 mM CaCl2 1M stock 1 mL

1 mM MgCl2 1M stock 1 mL 10 mM Glucose 180.16 g/M 1.8016 g 5 mM HEPES 238.31 g/M 1.1916 g pH buffered to 7.40 with 1M NaOH Solution was stored at 4 °C

Table IV: Preparation of internal solution for patch-clamp electrophysiology experiments

Internal (pipette) solution components

Final Volume = 40 mL (in ddH20)

Final Conc. Compound M.W. Weight/Vol

140 mM KCl 74.55 g/M 0.4175 g

1 mM MgCl2 1M stock 40 μL 5 mM EGTA 380.40 g/M 0.0761 g 10 mM HEPES 238.31 g/M 0.0953 g *5 mM ATP (Mg salt) 507.18 g/M 0.1140 g pH buffered to 8.0 with 1M KOH *Add ATP and buffer pH to 7.20 with 1M KOH Filter to 1.5 mL tubes, label and store at -80 °C

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Mississauga, ON). Briefly, equal amounts of protein lysate were incubated with components of a colorimetric assay kit for 15 min. Samples were optically analyzed at 595 nm following calibration with a standard curve of known protein concentration using a Beckman DU-640 spectrophotometer

(Beckman-Coulter, Fullerton, CA). Next, equal amounts of protein were subjected to SDS- polyacrylamide gel electrophoresis (7.5% gel for detection of hERG protein and 10% gel for detection of STX1A protein) for 1 h at 100 mV and then 1-1.5 h at 150 mV or until the loading dye reached the bottom of the gel. Next, proteins were electrophoretically transferred onto PVDF membranes at 100 mV for 1 h. The membranes were blocked with 5% non-fat dried milk in TBST and subsequently incubated with primary antibodies directed against the various proteins of interest (see below) for 2 h at room temperature, or at 4 °C overnight in 1% bovine serum albumin (BSA). Secondary antibodies used were horseradish peroxidase-conjugated donkey anti-mouse IgG or donkey anti-goat IgG

(Jackson Immunoresearch, West Grove, PA) and donkey anti-rabbit IgG (Santa Cruz Biotechnology,

Santa Cruz, CA). Secondary antibodies were used in the range from 1:20,000 – 1:40,000 and were diluted in 5% milk for 1 h at room temperature. The antibodies were detected with an ECL plus detection kit (GE Healthcare) or Western Lightning chemiluminescence agent (Perkin Elmer, Waltham,

MA) depending on the strength of the protein signal. Signals were exposed to X-ray films, which were developed using an automated photo-processor device (Kodak, Rochester, NY).

3.7 Antibodies

Several hERG primary antibodies were used for various experiments. For molecular biology experiments, WT hERG was detected using PA3-860, a polyclonal rabbit antibody directed against a

181 amino acid epitope of the hERG C-terminus, used at a 1:5000 dilution unless otherwise noted in the figure legends (ABR Affinity Bioreagents, Golden, CO).

Alternatively, several other anti-hERG antibodies were used for various experiments including: APC-

062 (Alomone, Jerusalem, Israel) corresponding to a 54 amino acid epitope and APC-016 (Alomone)

-58- corresponding to a 16 amino acid C-terminal epitope. A polyclonal rabbit antibody APC-109

(Alomone) directed against a 16 amino acid epitope located between the S1 and S2 transmembrane domains was also used as an external marker for hERG protein. In addition, two purified goat polyclonal antibodies were also utilized. hERG C-20 (sc-15968, Santa Cruz Biotechnology,) consists of a short epitope directed against the C-terminus, while hERG N-20 (sc-15966, Santa Cruz) consists of a short epitope directed against the N-terminus.

HA-hERG was detected during immunoblot analysis using polyclonal rabbit anti-HA antibody (Sigma

H3663) at a 1:5000 dilution and STX1A was detected using monoclonal mouse anti-STX1A antibody

(Sigma S0664) at a 1:5000 dilution unless otherwise noted by the figure legend. Anti-β-actin (Sigma

A5316) and α/β tubulin (Cell Signaling Technology 2148, Danvers, MA) were used as loading controls in concentrations ranging from 1:1000 – 1: 5000. All antibodies were prepared in TBST containing BSA according to the manufacturers’ instructions, were stored at 4 °C between uses, and were not used beyond 2 months after preparation.

3.8 Immunocytochemistry and confocal microscopy

Stably transfected hERG-HEK 293 cells without or with STX1A transfection were plated on sterile 18 mm glass coverslips for 24 h to similar confluency for immunofluorescent studies. Cells were fixed for

30 min with 2% paraformaldehyde (Sigma) and washed for 5 min with 25 mM NH4Cl/PBS and then 3 times (5 min each) with phosphate buffered saline (PBS). Cells were blocked and permeabilized for 1 h with 0.1% saponin blocking buffer containing, 5% normal goal serum (Jackson) and PBS. They were then incubated overnight at 4 °C with primary antibodies: rabbit anti-hERG (ABR, PA3-860) and mouse anti-STX1A (Sigma, monoclonal) at 1:250 dilution, followed by five 5-10-min washes with PBS and a 1 h incubation with secondary antibodies (anti-rabbit IgG labeled with fluorescent isothiocyanate (FITC) for hERG and anti-mouse IgG with tetra-methyl-rhodamine-isothiocyanate (TRITC) for STX1A).

Confocal microscopy was performed with a Zeiss LSM-510 system (Carl Zeiss Canada, Toronto, ON).

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TRITC (red) and FITC (green) were excited at 543 and 488 nm with HeNe- and Ar-Lasers respectively, emitting fluorescence at 585 and 505-530 nm. Confocal microscopy experiments were performed on the same day for both groups, with identical parameters used for all manipulations. Control experiments omitting primary antibodies and with non-transfected hERG-HEK293 cells revealed absent or very low-level background fluorescence.

3.9 In vitro binding studies

Forty-eight hours after transfection, cells were washed with PBS (pH 7.4) and then harvested in binding buffer (25 mM HEPES (pH 7.4) 100 mM KCl, 1.5% Triton X-100, 2 μM pepstatin A, 1 μg/ml leupeptin and 10 μg/ml aprotinin). The cells were lysed by sonication and insoluble materials were removed by centrifugation at 22,500 × g at 4 °C for 30 min. The detergent-extract (0.3 ml, 1.5-2.1

μg/ml protein) of cell lysate was incubated with GST (as a negative control) or GST-STX1A, GST-STX1A-

HABC, or GST-STX1A-H3 fusion proteins (all bound to glutathione-agarose beads, 500 pM protein at 4 °C for 2 h). The beads were then washed three times with binding buffer and eluted proteins were analyzed by SDS-PAGE. Protein was transferred to PVDF membranes, blotted with primary hERG antibody (APC-062, Alomone) and secondary goat anti-rabbit antibody, and detected using ECL Plus

(GE Healthcare).

3.10 Coimmunoprecipitation

HEK 293 cells grown on 100 mm plates were transfected with hERG (or HA-hERG WT or hERG truncation mutations) and STX1A were washed twice with ice-cold PBS following 48 h incubation.

Cells were then lysed with 1 mL lysis buffer added directly to the cell culture dish and incubated on ice for 25 min. Lysis buffer contained: 50 mM Tris-HCl pH 8.0, 150 mM NaCl, 1 mM CaCl2, protease inhibitor cocktail (Roche) and 1% Triton X-100. Cell lysates were then scraped into 1.5 mL Eppendorff tubes and incubated on ice for approximately 30 min. During this time, lysates were passed 10 times

-60- through a 20G 1½“ needle to break up the cell lysate. Samples were then centrifuged at 16,000 x g for

30 min to clear nuclei and debris. Supernants were retained and protein assay was performed as described in the Western blot section using a modified Bradford method. Next, 50 μL of 50% slurry of

Protein A Sepharose CL-4B resin (GE Healthcare) were added to each sample to pre-clear the lysates of nonspecific binding. These samples were incubated at 4 °C for 1 h and placed on a mixer, and then centrifuged at 16,000 x g for 20 min. Supernants were retained, and a sample was set aside as a cell lysate control. The remainder of the sample (at least 750 μg protein per group) was incubated with 50

μL of a 50% slurry of Protein A beads and 2 μL of antibody (either anti-STX1A or anti-HA/hERG antibodies). The mixtures were mixed gently for 2 h at 4 °C on a shaker. After binding the bead/antibody/protein complexes, cell lysates were washed 3-4 times with wash buffer containing 10 mM Tris-HCl pH 8.0, 150 mM NaCl and 0.1% Triton X-100 and centrifuged at 800 rpm for 2 min between washes. Supernants were discarded. During the final wash, 100 μL of beads and wash buffer were left in the 1.5 mL Eppendorff tubes, and 20 μL of 6X SDS loading buffer for a final concentration of 50 mM Tris-HCl pH 7.0, 5% glycerol, 1.67% SDS, 83 mM DTT and 0.0002% bromophenol blue.

Proteins were eluted by boiling the sample tubes for 5 min prior to loading for SDS-PAGE for immunoblotting analysis as outlined above. Membranes were immunoblotted for the putative interacting protein partner and precleared control lysates were immunoblotted to ensure the presence of both putative interacting partner proteins.

3.11 Isolation of endogenous protein including membrane protein

Wild-type C57BL/6 adult mouse hearts (Charles River, St. Constant, QC) were removed and cardiac membrane protein isolation was performed in a similar fashion as described previously (Pond et al.,

2000). Briefly, 4-6 mice were anesthetized and hearts were rapidly removed and frozen on liquid N2 for

20 min. All procedures were performed at 4 °C hereafter with protease inhibitors (Roche). Frozen hearts were smashed on ice to break cells and tissues were then thawed and homogenized in 10 mL of

TE buffer using a tissue homogenizer. TE buffer contained 10 mM Tris-HCl pH 7.4, 1 mM EDTA and

-61- protease inhibitors. Samples were then centrifuged at 1000 rpm for 10 min to clear nuclei and debris.

Supernants were kept in one tube and pellets were kept in another and resuspended and centrifuged again at 1000 rpm for 10 min. Both sets of supernants were then pooled and centrifuged at 100,000 x g for 10 min. These pellets were kept and resuspended in TE buffer containing 0.6 M KI and incubated on ice for 30 min. Incubation with high KI was necessary to destabilize contractile proteins which constitute the majority of the protein present in the cell lysates. Samples were then centrifuged at

40,000 x g for 10 min. Supernatants were removed and the pellet was resuspended in TE buffer to wash KI and then samples were centrifuged at 40,000 x g for 10 min. This was repeated to completely wash KI from the lysate. The final pellet was resuspended in TE buffer containing 2% Triton X-100 and incubated on ice for 1 h to solubilize the membrane proteins. A final centrifugation was carried out at

17,400 x g for 30 min to precipitate insoluble material. Supernants were then frozen at -20 °C until use and protein assay was carried out to measure protein concentration as described above.

3.12 Immunoblot quantification and statistical analysis

Digital images were produced as *.tif files in grayscale format from X-ray films of Western blots and were quantified using Image J (NIH). Quantification of immunoblot band optical density is calculated by determining the sum of the pixel intensity for each band subtracted by the average background signal. Average pixel values were recorded and analyzed in Microcal Origin 6.0TM (OriginLab

Corporation, Northampton, MA). Quantifications are presented as arbitrary units normalized to loading or alternative protein expression markers.

For all figures, data points represent mean ± S.E.M. and “n” is the number of experiments per group.

Origin was used to analyze data and fit exponential functions to raw current traces. An unpaired

Student’s t test was used to compare control values to experimental groups. A p < 0.05 was considered statistically significant. In experiments comparing more than 2 groups, one-way analysis

-62- of variance (ANOVA) for repeated measures was calculated. Statistical significance is denoted in the figures by * (p < 0.05), ** (p < 0.01) and *** (p <0.001).

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Chapter 4: Results

4.1 Determining an ideal system for electrophysiological assessment of hERG channels

Previous reports characterizing the expression of hERG currents have employed the use of oocyte expression and transient transfection techniques. We chose to examine hERG currents recorded at room temperature (20-22 °C) following transient transfection in tsA-201 cells which are derived from human embryonic kidney cells as described in the materials and methods section (Margolskee et al.,

1993). Fig. 11 illustrates the properties of hERG currents recorded in this transient transfection system.

Fig. 11 A shows typical original current recordings from tsA-201 cells transfected with hERG (upper traces) and tsA-201 cells without hERG transfection (lower traces). Currents were elicited from a holding potential of -80 mV, and then subjecting them to 3-s depolarizations from -80 to +60 mV in

+10 mV increments. Immediately following depolarization, cells were repolarized to -60 mV for 3 s before returning to the holding potential of -80 mV. Outward K+ currents were activated at potentials positive to -40 mV reaching their peak conductance at +20 mV. At progressively more positive potentials, hERG currents were reduced as a result of C-type inactivation (Sanguinetti et al., 1995) as clearly illustrated by inward rectification of the upper set of current traces. Untransfected tsA-201 cells possess background outward K+ current which increases with more positive depolarizations. Fig. 11 B illustrates the I-V relationship of populations of hERG and endogenous currents measured at the end of the 3-s depolarizing pulses. hERG-transfected cells reached a maximum current amplitude of 136.1

± 7.6 pA/pF at +20 mV (n = 18 cells) vs. untransfected cells which reached a maximum current amplitude of 51.8 ± 3.8 pA/pF at +60 mV (n = 5 cells). Repolarization facilitated the recording of hERG tail currents which increased in current amplitude following progressively more positive depolarizations. Untransfected tsA-201 cells do not produce tail currents and this clearly illustrates the lack of endogenous hERG channel expression in this cell type. Fig. 11 C shows the I-V relationship for hERG tail currents measured at the beginning of the 3-s repolarizing pulse. Following normalization of

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Figure 11. Expression of hERG WT cDNA in tsA-201 cells.

Transient transfection of tsA-201 cells with hERG cDNA facilitated the recording of single-cell currents following step depolarizations from -80 mV to +60 mV, followed by repolarizing pulses to -60 mV. A) A family of typical hERG currents elicited from transiently transfected tsA-201 cells (upper traces). hERG currents were elicited at potentials positive to -40 mV reaching a peak amplitude at +20 mV and undergoing inward rectification at progressively more positive potentials. B) Untransfected tsA-201 cells do not possess hERG current, but do possess a measurable amount of outward potassium current, which increases with progressively more depolarized potentials (lower traces). B) Current-voltage relationship for hERG-transfected tsA-201 cells and untransfected cells at the end of a 3-s test pulse. hERG-transfected cells (n = 18 cells) reach their peak current amplitude at +20 mV, while untransfected cells (n = 5 cells) reach peak current amplitude at +60 mV. C) Current- voltage relationship for normalized tail currents for hERG-transfected cells (n = 18 cells) taken at the beginning of 3-s repolarizing pulses was fit to a Boltzmann’s equation to determine the V1/2 of steady-state activation and slope factor.

-65- the maximum tail current amplitude, the voltage-dependence of channel activation curve was derived by fitting the data points to a Boltzmann function:

/ where I is the relative tail current, V1/2 is the voltage required for half activation of current, Vt is the test potential, and k is the slope factor (V1/2 = -9.1 ± 1.1 mV; k = 8.5 ± 0.3 mV; n = 18). The steady-state activation curve shows that the activation threshold for hERG transfected tsA-201 cells is about -40 mV with full activation occurring at potentials positive to +20 mV. The average current density at +60 mV was 114.6 ± 16.3 pA/pF (n = 18 cells).

Due to a significant amount of endogenous current present in tsA-201 cells, we obtained HEK 293 cells which were stably transfected with hERG. Fig. 12 A shows currents elicited from a single stably- transfected hERG-HEK 293 cell (upper traces). To characterize any underlying endogenous currents affecting the expression of hERG currents, we used the high-affinity hERG channel blocker E-4031

(Sanguinetti and Jurkiewicz, 1990; Jurkiewicz and Sanguinetti, 1993). E-4031 is a methanesulfonanilide drug which has been reported to block hERG currents with high affinity. It was previously reported that the dose-response curve for E-4031 for stably transfected hERG-HEK 293 cells had an EC50 of 7.7 nM and a Hill coefficient of 1.0 (Zhou et al., 1998b). In Fig. 12 A (middle traces), hERG currents were almost completely blocked with 300 nM of E-4031 perfusion for 15 min resulting in a substantial reduction in hERG current amplitude, particularly in the outward hERG tail currents. This figure illustrates the expression of a measurable amount of endogenous outward current, albeit substantially smaller than those measured from tsA-201 cells. Fig. 12 A (lower traces) illustrates that block due to E-4031 drug binding is reversible and peak current amplitude is partially recovered following 1-h wash-out. Fig. 12 B depicts the time-course of E-4031 block and wash-out as measured by current amplitude at the end of a 3-s depolarizing pulse to 0 mV occurring every 30 s. E-4031 reduced peak hERG current at 0 mV to 16% of baseline levels, and 1-h wash-out successfully recovered

41% of the blocked current amplitude. Fig. 12 C shows the hERG-HEK 293 I-V relationship for 6 cells

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Figure 12. Whole-cell currents elicited in stably transfected hERG-HEK 293 cells

Whole-cell hERG currents were elicited from hERG-HEK 293 cells by performing 3-s step depolarizations from -80 to +60 mV followed by repolarization for 3 s to -60 mV. A) Typical hERG-HEK 293 current traces - peak amplitudes occur at ~0 mV and undergo rectification (upper traces). This single cell was perfused with 300 nM E- 4031 for 15 min, resulting in a large reduction in hERG currents (as evaluated by peak tail current amplitude) revealing the contribution of endogenous outward current (middle traces). This hERG-HEK 293 cell was washed for 1 h to remove E-4031 which resulted in the partial recovery of hERG channel current. B) Time course for hERG channel block with E-4031 and washout. E-4031 was perfused for 15 min from t = 0 resulting in a reduction of current to 16% of baseline levels. After washing for 1 h, there was a 41% restoration of hERG current amplitude. C) I-V relationship for hERG-HEK 293 cells without and with 300 nM E-4031 perfusion for 15 min (n = 6 cells). hERG currents were significantly reduced by E-4031, revealing an endogenous outward current which increases in current amplitude with progressively more depolarized test potentials. D) Tail current amplitude reveals that hERG currents were almost completely blocked, therefore verifying the assumption that currents observed in C) in the presence of E-4031 are mostly due to the contribution of endogenous outward current.

-67- before and during 300 nM E-4031 block. Baseline hERG currents achieved a peak current amplitude of

53.3 ± 8.9 pA/pF at 0 mV, while current amplitude was reduced to 5.3 ± 1.6pA/pF at 0 mV following 15 min of 300 nM E-4031 perfusion. Currents at +60 mV for E-4031 were 7.1 ± 2.0 pA/pF, and this gives an approximation for the amount of endogenous current present in hERG-HEK 293 cells. Compared to tsA-201 transient transfection, stably transfected hERG cells reach their peak current amplitude at 0 mV (as opposed to +20 mV) and undergo a greater degree of rectification (hERG-HEK 293: 26.9% of peak current at + 60 mV, n = 6 cells vs. tsA-201 cells: 49.9% of peak current amplitude at +60 mV, n =

18 cells, see Fig. 11 B). These differences may be attributed to the significant contribution of endogenous outward currents in naïve tsA-201 cells, thus supporting our decision to use hERG-HEK

293 cells for further electrophysiological assessment of the channel. To exclude the possibility that not all hERG channels were blocked by E-4031, hERG tail current amplitudes were measured at the beginning of the 3-s repolarizing pulse to measure hERG tail current, which does not possess any endogenous current. Fig. 12 D illustrates the I-V relationship for hERG tail currents at +60 mV measuring 61.6 ± 4.7 pA/pF (hERG-HEK 293 control) compared to E-4031 treatment of the same cells,

2.3 ± 1.6 pA/pF (hERG-HEK 293 + 300 nM E-4031, n = 6 cells). Although a minute amount of measurable current is still present, it is extremely impaired compared to control and is also significantly smaller than currents recorded during depolarizing pulses in E-4031-treated cells. This illustrates the effectiveness of E-4031 in blocking hERG current. Based on the aforementioned observations, we continued to utilize the hERG-HEK 293 model system for the electrophysiological characterization of a potential hERG-STX1A interaction. (Preliminary experiments were carried out using tsA-201 cells, and this data is included in Appendix 1).

4.2 STX1A significantly reduces hERG current amplitude

Endogenous expression of STX1A in isolated rat ventricular myocytes has recently been demonstrated

+ as well as the STX1A-mediated modulation of cardiac ATP-sensitive K currents and KV4.2 channels

(Kang et al., 2004; Yamakawa et al., 2007). Therefore, we also reasoned that STX1A may interact with

-68- other cardiac ion channels such as hERG. Making them ideal candidates for SNARE protein interaction is the ability for hERG channels to enthusiastically interact with auxiliary subunits, which alter their expression and gating.

In order to electrophysiologically assess the potential hERG-STX1A interaction, hERG-HEK 293 cells were transfected without or with STX1A and GFP. Successful transfection was visualized by green fluorescence. Fig. 13 A used the same voltage protocol described in Fig. 12 to elicit hERG currents from stably-transfected control hERG-HEK 293 cells (upper traces) and hERG-HEK 293 cells transiently transfected with STX1A (bottom traces). Coexpression of hERG with STX1A dramatically reduced current amplitude with no change in C-type inactivation. Fig. 13 B shows the I-V relationship for hERG and hERG + STX1A. Current amplitude was measured at the end of the 3-s depolarizing pulses. Peak current amplitude was reduced from 71.3 ± 4.7 pA/pF (hERG, n = 35 cells) to 26.7 ± 2.9 pA/pF at +10 mV (hERG + STX1A, n = 24 cells, p < 0.001). The inset illustrates that despite a large difference in current amplitude, hERG and hERG + STX1A groups undergo C-type inactivation to a similar extent as illustrated by this normalized I-V relationship. Fig. 13 C highlights the tail currents elicited by the 3-s repolarizing pulse to -60 mV. hERG + STX1A tail currents are also severely reduced. Fig. 13 D shows the I-V relationship for tail currents measured at the beginning of the 3-s repolarizing pulse. Peak tail currents measured at +60 mV were reduced from 66.2± 3.4 pA/pF (hERG, n=35) to 32.5 ± 2.7 pA/pF

(hERG + STX1A, n=24; p<0.001). These I-V plots were fit with a Boltzmann equation which revealed that STX1Ahad no significant influence on the midpoint of current activation (V1/2: -16.6 ± 0.9 mV hERG vs. -13.6 ± 1.8 mV hERG + STX1A) or slope (k: 9.0 ± 0.2 hERG vs. 9.7 ± 0.3 hERG + STX1A). The inset shows the normalized I-V relationship for the tail currents, which are virtually superimposable.

4.3 STX1A affects steady-state inactivation but not gating kinetics

Time constants for hERG channel activation were determined using two methods as described previously (Sanguinetti et al., 1995; Zhou et al., 1998b). At test potentials ranging from -40 to 0 mV,

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Figure 13. STX1A significantly impairs hERG current amplitude

Stably transfected hERG-HEK 293 cells were used to assess the electrophysiological interaction of hERG and STX1A. A) Whole-cell hERG currents (upper traces) were significantly reduced when STX1A was transiently transfected (lower traces). The blue box indicates the currents elicited by step-depolarizations from -80 mV. B) I- V relationship measured for hERG-HEK 293 cells (n = 35 cells) and hERG-HEK 293 cells + STX1A (n = 24 cells). STX1A significantly reduces hERG current amplitude without affecting C-type inactivation (see inset, normalized I-V relationship). C) hERG-HEK 293 tail currents are highlighted without and with transient STX1A transfection. D) I-V relationship for tail currents demonstrates hERG tail currents (n = 35 cells) are significantly reduced by STX1A (n = 24 cells). Inset shows the normalized I-V relationship for tail currents demonstrating that there was no significant shift in the midpoint of steady-state current activation or its slope factor.

-70- activation time constants were obtained from the rising phase of continuous currents measured following 3-s depolarizing pulses. These continuous currents were fit with a mono-exponential function in order to obtain activation time constants. Original current traces are shown in Fig. 14 A with the mono-exponential fit superimposed in red for the various test potentials. At more positive potentials, activation time constants were obtained by recording an envelope of tail currents at various test potentials (0, +20, +40 and +60 mV). Envelope currents were produced by depolarizing cells to a given test potential for increasing durations and then repolarizing them with a pulse to -120 mV to produce tail current. An original current recording for envelope currents produced at 0 mV is illustrated on Fig. 14 B. Peak tail currents from these recordings were extracted and then fit to a mono-exponential function in order to obtain the activation time constant. Fig. 14 C illustrates data obtained from the same cell as Fig. 14 A and B measuring envelope currents at the four test potentials.

Although obtaining activation time constants from continuous currents is less time consuming, it is not necessarily the most precise method at more positive potentials as the effect of C-type inactivation becomes more prevalent. Fig. 14 C shows that time constants for activation using the same cell and the two methods produce slightly different results for activation time constants. Using the continuous current method, the activation time constant was 386 ms, whereas the time constant obtained from the envelope currents was 269 ms. Despite these different values, the mean time constants for activation at 0 mV were not significantly different between the two groups (see Fig. 14

E). Generally, the continuous current method was acceptable for measuring currents at potentials below 0 mV, while the envelopes method was more suitable for obtaining time constants at more positive potentials because of the strong influence of hERG channel inactivation at progressively more positive test potentials. Fig. 14 E summarizes all of the activation time constant data for hERG-HEK 293 cells without and with STX1A. Activation time constants were obtained from continuous current recordings (closed shapes) from -40 to 0 mV and were not significantly different at any test potential between hERG (n = 35 cells) and hERG + STX1A (n=24 cells). Additionally, there was no significant difference at any potential from 0 to +60 mV between hERG (n = 22-28 cells) and hERG + STX1A (n =

12-16 cells) as obtained from envelope currents (open shapes).

-71-

Figure 14. STX1A has no effect on hERG channel activation Two methods were used for obtaining hERG activation time constants. A) The rising phase of continuous currents from test potentials ( -40 to 0 mV) were fit to a mono-exponential function (fits in red). B) At positive potentials, tail current envelopes were elicited and then plotted D) to fit envelopes to a mono-exponential function. C) Data obtained for the same cell using both methods at 0 mV. At positive voltages, the influence of hERG inactivation affects continuous current traces and the envelopes method is more accurate. E) Summarized data for activation time constants using continuous currents (closed symbols, hERG: black squares n = 35 cells; hERG + STX1A: red circles n = 24 cells) and using envelope currents (open symbols, hERG n = 22-28 cells; hERG + STX1A n = 12-16 cells). There was no significant difference between hERG and hERG + STX1A groups at any test potential. There was also no difference in mean activation time constant at 0 mV for the two methods utilized.

-72-

The fully-activated I-V plot for hERG was obtained by depolarizing cells to +60 mV for 1 s followed by repolarization to voltages ranging from -120 to +60 mV. Representative current traces from a control hERG-HEK 293 cell are shown (Fig. 15 A, lower panel). At the most negative potentials, the currents are inward, and as channels are repolarized to more positive potentials, reversal occurs at approximately -

85 mV. hERG channels produce outward current reaching a peak potential at -20 mV, and then rectify.

Current amplitude was measured at the start of the 5-s repolarizing pulse and plotted in Fig. 15 B.

Peak current amplitude for hERG cells (112.7 ± 11.8 pA/pF at -20 mV, n = 14 cells) was significantly larger than hERG + STX1A (50.1 ± 9.6 pA/pF at -20 mV, n = 8 cells; p < 0.001). The normalized I-V relationship is shown in the inset. Time constants for hERG channel deactivation were obtained by fitting tail current decay to a double exponential function. Fig. 13 C illustrates five representative tail currents fitted with the function resulting in a fast and a slow time component of hERG channel deactivation. These time constants are voltage-dependent as shown by Fig. 15 D which summarizes the deactivation time constants for hERG (n = 14 cells) and hERG + STX1A (n = 8 cells). There was no significant difference between hERG and hERG + STX1A for either the fast (open shapes) or slow time constants of deactivation (closed shapes). The relative contribution to current amplitude of the fast and slow current decays is shown in Fig. 15 E. Data represents the mean contribution of fast current decay relative to the total current decay (AFast / AFast + ASlow). At negative potentials, the amplitude of the fast component is much larger, and the amplitude of the slow component becomes increasing predominant with more positive test potentials. There was no significant difference in the relative contribution of the fast or slow components of current amplitude decay between hERG (n = 14 cells) and hERG + STX1A (n = 8 cells).

The kinetics of hERG channel fast inactivation and recovery from inactivation were studied independently using protocols illustrated in Fig. 16. Firstly, hERG channel recovery was calculated from currents recorded with a two-pulse protocol. To evaluate hERG channel recovery from inactivation, currents were elicited following a 200 ms depolarizing pulse to +60 mV to activate and then inactive hERG channels, followed by step repolarizations from -100 to -20 mV (Fig. 16 A).

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Figure 15. STX1A does not affect hERG channel deactivation

A fully activated I-V plot for hERG tail currents was obtained by depolarizing cells for 1 s at + 60 mV followed by repolarizing cells in +10 mV increments from -140 to +60 mV and then recording a tail current. A) A typical family of current traces from a hERG-HEK 293 cell. Current amplitude was measured immediately following the depolarizing pulse in order to obtain the I-V plot in B). hERG-HEK 293 tail currents have a reversal potential of approximately -85 mV and reach a peak amplitude at approximately -20 mV before rectifying (hERG, n = 14 cells). Transient transfection with STX1A does not affect any of these parameters, however, does cause a significant reduction in current amplitude (hERG + STX1A, n = 8 cells). Inset illustrates the normalized I-V relationship. C) Decaying tail currents are fitted with a double exponential function in order to obtain two time constants for hERG channel deactivation (representative traces are shown). D) Summarized mean time constant data for hERG (black squares, n = 14 cells) and hERG + STX1A (red circles, n = 8 cells) deactivation time constants. STX1A had no affect on either the fast (open symbols) or slow (closed symbols) time constants of channel deactivation. E) Summarized data illustrating the relative contribution to current amplitude of the fast component of current decay relative to total current. There was no significant difference between hERG and hERG + STX1A at any test potential.

-74-

Figure 16. STX1A does not affect fast inactivation or recovery from inactivation A) Time constants for recovery from inactivation were obtained following a brief depolarizing pulse to +60 mV followed by repolarizing pulses from -100 to -20 mV. The resulting currents were fit with a mono exponential function in order to obtain time constants (see B) for example). C) Time constants for hERG fast inactivation were obtained using the triple-pulse protocol illustrated (upper panel). Families of current traces were obtained (lower panel) and fit to a mono exponential function as in D) for test potentials ranging from -20 to +60 mV. E) Summarized data for mean time constants for recovery from inactivation (closed symbols) and fast inactivation (open symbols). There was no significant difference in the recovery time constants between hERG (black squares, n = 11 cells) and hERG + STX1A (red circles, n = 6 cells) at any test potential. Also, there was no significant different in the time constants for fast inactivation between hERG (n = 11 cells) and hERG + STX1A (n = 5 cells).

-75-

A representative current trace of a control hERG-HEK 293 cell is shown (lower panel). Individual current traces were fit to a mono-exponential function as in Fig. 16 B in order to obtain a time constant for hERG channel recovery from inactivation. Fits were made to both inward and outward currents produced at the immediate start of the repolarizing pulse. There was no difference in recovery from inactivation between hERG (n = 11 cells) and hERG + STX1A (n = 6 cells) as summarized in Fig. 16 E

(closed symbols). hERG channel fast inactivation was measured from recordings generated from the three-pulse protocol shown in Fig. 16 C. Cells were first depolarized for 200 ms to +60 ms to activate and inactivate hERG channels, and were then repolarized to -100 mV rapidly for 10 ms to recover

channels without significant deactivation (deactivation τFast at -100 mV = 120 ms). This was followed by step depolarizations from -20 to +60 mV to elicit currents in order to record inactivation hERG currents (bottom panel). These rapidly inactivating currents were fitted with a mono-exponential function in order to obtain a time constant for hERG channel inactivation (Fig. 16 D for examples).

Mean inactivation time constant data is summarized in Fig. 16 E (open symbols). There was no difference between hERG (n = 11 cells) and hERG + STX1A (n = 5 cells) at any test potential.

Although we did not observe any STX1A-mediated changes in the time constants of hERG channel activation, deactivation, inactivation or recovery, we hypothesized that STX1A may still affect hERG channel gating. The intrinsic property of hERG channel inward rectification is a reflection of reduced channel availability at increasingly depolarized test potentials. The steady-state inactivation parameter compares channel availability between depolarized and hyperpolarized potentials. A reduction in hERG current amplitude could be explained by modulation of the steady-state inactivation parameter during interaction with STX1A. Steady-state inactivation was measured using the triple-pulse protocol shown in Fig. 17 A. This model of steady-state inactivation may not fully reflect the complex nature of hERG channel gating; however, it has been well established as an appropriate and accurate method for the approximation of steady-state inactivation (Smith et al.,

1996; Nakajima et al., 1998; Kiehn et al., 1999). Steady-state inactivation was investigated by first depolarizing single cells for 2 s to a potential of +60 mV, followed by variable short repolarizations

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Figure 17. STX1A induces a hyperpolarizing shift in the midpoint of steady-state inactivation

A) Steady-state inactivation was assessed using a triple-pulse protocol consisting of a 2-s depolarizing pulse to +60 mV followed by step repolarizations from -140 to +20 mV (20 ms, 10 mV steps). Outward currents were measured during the immediate onset of a subsequent 500 ms depolarizing pulse to +20 mV. B) Peak currents elicited during the final depolarizing pulse were plotted versus the preceding test potential and normalized. Steady-state inactivation curves were constructed for hERG (black squares, n = 15 cells) and hERG + STX1A (red circles, n = 10 cells) by fitting data to the Boltzmann function. STX1A induced a significant hyperpolarizing shift in the midpoint of steady-state inactivation while having no effect on slope factor.

-77- ranging from -140 to +20 mV (20 ms, +10 mV increments) so that channels could be recovered from inactivation. Finally, a depolarization to +20 mV for an additional 500 ms permitted the recording of an outward current (Fig. 17 A; lower panel). Peak currents measured within the first 2 ms of the final depolarizing pulse were plotted versus the preceding test potential (Fig. 17 B). These currents were normalized and then fitted to the Boltzmann function. STX1A induced a hyperpolarizing shift in the midpoint (V1/2) of hERG steady-state inactivation (hERG, n = 15 cells, -34.4 ± 2.3 mV vs. hERG + STX1A, n

= 10 cells, -47.7 ± 4.1 mV; p < 0.01). STX1A did not affect the slope (k) of steady-state inactivation (20.6

± 0.6 mV hERG vs. 18.6 ± 0.5 mV hERG + STX1A). This result implies that the STX1A-mediated impairment of hERG current amplitude may be partially explained by a decrease in the proportion of available hERG channels. Alternatively, this result implies that in the presence of STX1A, there is an increased likelihood that hERG channels would transition to the inactivated state at more negative potentials. However, this slight hyperpolarizing shift in steady-state inactivation could not possibly fully account for the overwhelming reduction in hERG current amplitude observed in Fig. 13 B.

4.4 STX1A-imparied hERG current amplitude is partially restored by E-4031

Several studies have demonstrated the ability of the high-affinity hERG pore-blocking compound E-

4031 to rescue the trafficking and current density of several mutant hERG proteins (Zhou et al., 1998a;

Zhou et al., 1999; Robertson and January, 2006). We decided to test the hypothesis that E-4031 may be able to increase the current density of STX1A-impaired hERG channels. hERG-HEK 293 cells without and with STX1A transient transfection were incubated without or with 5 μM E-4031 for 24 h prior to use for electrophysiological experiments. Dishes containing cells treated with E-4031 were rinsed with fresh medium for 1 h prior to recording. Following 24-h incubation with E-4031, hERG + STX1A peak current amplitude was significantly restored (Fig. 18 A) to 50.7 ± 4.7 pA/pF (hERG + STX1A + E-4031, n

= 19 cells) vs. hERG + STX1A (26.7 ± 2.9 pA/pF, n = 24 cells) (p < 0.001); and was not significantly different from hERG + E-4031 control (54.7 ± 3.1 pA/pF, n = 41 cells). E-4031 incubation also partially rescued tail currents (Fig. 18 B) from 32.5 ± 2.7 pA/pF (hERG + STX1A, n = 24 cells) to 46.0 ± 2.3 pA/pF

-78-

Figure 18. E-4031 can partially rescue STX1A-imparied hERG current amplitude hERG-HEK 293 cells without or with STX1A transient transfection were incubated without or with 5 μM E-4031 for 24 h and then used for electrophysiological assessment. A) I-V relationship for hERG control (n = 35 cells), hERG + STX1A (n = 24 cells), hERG + E-4031 (n = 41 cells) and hERG + STX1A + E-4031 (n = 19 cells). E-4031 incubation significantly increased hERG + STX1A current amplitude and was not significantly different from hERG + E-4031 control. B) I-V relationship for tail currents obtained for the same groups of cells as in A). E-4031 significantly increased hERG + STX1A current amplitude which was also not significantly different from hERG + E- 4031 control. C) The STX1A-mediated hyperpolarizing shift in steady-state inactivation was not rescued following 24-h incubation with E-4031. Furthermore, the modest hyperpolarizing shift in the V1/2 of hERG + E- 4031 relative to hERG control was not statistically significant. (hERG control n = 15; hERG + STX1A n = 10 cells; hERG + E-4031 n = 17 cells; hERG + STX1A + E-4031 n = 8 cells).

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(hERG + STX1A + E-4031, n = 19 cells; p < 0.001) which also was not significantly different from hERG +

E-4031 control (53.5 ± 2.5 pA/pF, n = 41 cells). E-4031 can partially rescue the reduction in current amplitude caused by STX1A. In addition, steady-state inactivation was investigated in order to ascertain whether E-4031 could cause any change in the hyperpolarizing shift caused by STX1A (Fig.

18 C). Incubation with E-4031 for 24 h had no effect on the midpoint of steady-state inactivation of hERG expressing cells cotransfected with STX1A (hERG + STX1A, n = 10 cells, -46.4 ± 3.7 mV vs. hERG +

STX1A + E-4031, n = 8 cells, -46.4 ± 3.7 mV). The modest hyperpolarizing shift in the midpoint of steady-state inactivation in E-4031-treated hERG cells was not statistically significant from hERG cells alone (hERG, n = 15 cells, -34.4 ± 2.3 mV vs. hERG + E-4031, n = 17 cells, -40.4 ± 2.6 mV). Overall, these results suggest that the E-4031-mediated rescue of hERG + STX1A current density is independent of the functional interaction between hERG and STX1A proteins.

4.5 Colocalization of hERG and STX1A

Coexpression of hERG and STX1A resulted in a significant reduction in current amplitude (see Fig. 13

B). The hyperpolarizing shift in steady-state inactivation described in Fig. 17 could not fully account for this reduction in current, and no other changes in the kinetics of activation, deactivation, recovery and inactivation were detected. Therefore, we hypothesized that a reduction in the number of mature, functional hERG channels at the plasma membrane may account for the overall reduced current. Alternatively, alterations in single-channel conductances could account for changes in total current amplitude; however, single-channel recordings were not attempted. Additionally, a decrease in the open probability of hERG channels at a given test potential is possible because the parameters studied (activation, deactivation, inactivation and recovery) were only approximations of the true complex gating schemes by which hERG channels operate. Unfortunately, this requires highly complex modeling and interpretation of single-channel recordings and this was not feasible nor

-80- considered a high enough priority for this study. We did, however, consider our approximations to be adequate and do not believe that changes in hERG channel open probability occurred.

In order to study the cellular localization of the two proteins of interest, we performed immunocytochemistry and confocal microscopy. hERG-HEK 293 cells were plated on 18 mm glass cover slides without and with transient STX1A transfection. Cells were blocked, permeabilized and then incubated overnight with polyclonal rabbit anti-hERG antibody and monoclonal mouse anti-

STX1A antibody. Confocal microscopy revealed diffuse periplasmic and robust plasma membrane expression of hERG protein in hERG-HEK 293 cells (Fig. 19 A). The upper left panel shows a single cell being excited at 488 nm causing FITC fluorescence indicating hERG protein localization. The upper right panel shows the same cells undergoing excitation at 543 nm, and no signal is present as STX1A is absent. The bottom left panel shows a differential interference contrast (DIC) image of the cell being studied, with the bottom right panel showing a merge of the three panels. Fig. 19 B shows a single hERG-HEK 293 cell transiently transfected with STX1A. hERG channel expression, as seen in the upper left panel, is predominantly periplasmic with a large reduction in cell-surface expression as seen in the control image. Additionally, STX1A expression is also predominantly periplasmic with a lesser degree on the plasma membrane. The bottom right panel illustrates a high degree of colocalization between hERG and STX1A, particularly in highly dense intracellular compartments. These results suggest that

STX1A inhibits the surface expression of hERG while sequestering the protein to a yet to be determined periplasmic location.

4.6 STX1A impairs hERG protein maturation

Western blot analysis of hERG-HEK 293 cells revealed the expression of two bands, corresponding to an ER-resident core-glycosylated 135 kDa immature protein, and the second corresponding to the mature plasma membrane-associated 155 kDa protein (Fig. 20). During protein synthesis and folding, hERG channels undergo core glycosylation in the ER and must complete complex N-linked

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Figure 19. Colocalization of hERG and STX1A hERG-HEK 293 cells without and with STX1A transfection were grown on glass cover-slides, permeabilized and then incubated with anti-hERG (Affinity BioReagents, PA3-860, 1:250) and anti-STX1A (Sigma, S0664, 1:250) primary antibodies followed by FITC (green, hERG) or TRITC (red, STX1A) secondary antibodies. Confocal immunofluorescence reveals the expression of hERG and STX1A proteins. A) Top left panel shows that hERG protein localization is periplasmic and plasma membrane in a representative single hERG-HEK 293 cell. Top right panel shows that there is no STX1A expression in hERG-HEK 293 cells and bottom left panel is a differential interference contrast (DIC) image. Bottom right panel is a merge of the other 3 panels. B) Intracellular hERG localization in a representative single hERG-HEK 293 cell transiently transfected with STX1A has predominantly periplasmic subcellular localization with a reduced plasma membrane expression pattern (upper left panel). STX1A expression is strongly periplasmic and STX1A colocalizes with hERG protein in distinct highly dense intracellular compartments (upper right panel and merge image).

-82- glycosylation before exiting the ER-Golgi network. Glycoprotein addition promotes association with

ER resident chaperones, promotes correct folding and oligomeric assembly, prevents degradation and supports quality control (Hammond and Helenius, 1995; Petrecca et al., 1999). Cell lysates were obtained 48 h after transfection and equal amounts of protein were loaded in each lane of a 7.5% polyacrylamide gel. Fig. 20 shows that the expression of the 155 kDa mature hERG band in hERG-HEK

293 cells is weaker than the immature 135 kDa band, and following transient STX1A transfection, the expression of mature hERG protein is markedly reduced. In order to verify the N-linked glycosylation characteristic of the 155 kDa band, the two groups were incubated with 10 μg/mL tunicamycin for 48 h. Tunicamycin is an inhibitor of glycoprotein synthesis, and completely inhibits the N-linked glycosylation pathway while promoting the ER accumulation of hERG protein. Surprisingly, tunicamycin hERG-HEK 293 cells have been shown to produce recordable current, thereby proving that N-linked glycosylation is not required for assembly and trafficking of functional hERG channels

(Gong et al., 2002). Lanes 3 and 4 indicate that tunicamycin incubation resulted in a complete loss of

155 kDa mature hERG protein expression, while simultaneously shifting the immature hERG protein band to 132 kDa. This shift occurs because tunicamycin prevents hERG protein from undergo core- glycosylation after translation (Gong et al., 2002).

In order to further characterize the effect of STX1A on hERG channel maturation, naïve HEK 293 cells were transiently transfected with hemagglutinin (HA)-tagged hERG WT (HA-hERG) without and with

STX1A. HA-hERG has been shown to have identical properties to WT hERG (Akhavan et al., 2003).

Additionally, the quality and reproducibility of Western blots can be assured, as the specificity of this antibody is much greater than any of the commercially available hERG antibodies that I tested. Fig. 21

A shows HA-hERG protein expression without and with increasing cotransfection with STX1A. HA- hERG channels clearly yield 2 bands at 135 and 155 kDa, and additionally, separation of these bands allows for further analysis of mature vs. immature protein. As the ratio of STX1A coexpression is increased, the mature 155 kDa HA-hERG band decreases until it is nearly absent at 5.0 μg STX1A, while there is no appreciable change in immature core glycosylated HA-hERG at 135 kDa. Fig. 21 A (bottom

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Figure 20. Western blot analysis of hERG expression reveals two distinct bands hERG-HEK 293 cell lysates were harvested without or with STX1A transient transfection and detected with an anti-hERG primary antibody (ABR PA3-860, 1:5000) directed toward the C-terminus and corresponding to a 181 amino acid epitope. Immunoblotting reveals the expression of two distinct protein bands at 135 and 155 kDa (lane 1). The more pronounced 135 kDa band corresponds to an immature, core-glycosylated hERG protein, while the 155 kDa band is the mature complex-glycosylated form, having undergone complex glycosylation. STX1A reduces the expression of 155 kDa mature hERG protein (lane 2). 24 h treatment with tunicamycin, an inhibitor of N-linked glycosylation yields only 1 visible band for both groups, which has been shifted to 132 kDa (lanes 3 and 4) due to loss of core-glycosylation.

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Figure 21. STX1A reduces mature HA-hERG protein expression in a dose-dependent manner

Naïve HEK 293 cells were transfected with HA-hERG without or with increasing doses of STX1A. A) Western blot shows that HA-hERG expression yields two distinct bands at 135 and 155 kDa, which are easily distinguished from one another. Mature HA-hERG expression at 155 kDa decreases with increasing STX1A cotransfection level (lower panel). HEK 293 cells do not contain endogenous HA-hERG or STX1A (lane 8). B) Densitometric analysis of mature HA-hERG/ total HA-hERG protein expression vs. STX1A transfection dose (n = 3 experiments). Mature HA-hERG protein expression is related to STX1A expression, which is plotted on the right y-axis as a measure of arbitrary densitometric units.

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panel) illustrates the increasing expression of STX1A protein corresponding to increases in total cDNA transfection, which is detected as a 35 kDa band. STX1A expression could be seen at all transfection levels; however, lower transfection levels including 0.01 μg required prolonged exposure. Naïve HEK

293 cells do not possess any endogenous HA-hERG or STX1A (lane 8). Fig. 21 B illustrates the densitometric analysis of the HA-hERG and STX1A expression levels from 3 different sets of transfections. HA-hERG expression is plotted as the ratio of complex glycosylated protein to total HA- hERG protein, as this is the most common method of quantifying mature HA-hERG protein. This ratio was plotted versus STX1A transfection concentration (n = 3 experiments). Mature HA-hERG channel expression at the plasma membrane decreased from 0.30 ± 0.04 (no STX1A cotransfection) to 0.22 ±

0.05 with 0.5 μg STX1A, to 0.07 ± 0.03 with 5.0 μg STX1A. Expression of STX1A protein was quantified in arbitrary densitometric values (right y-axis) and plotted versus transfection amount. STX1A protein expression was linearly related to transfection amount. Overall, this result demonstrates that mature

HA-hERG protein expression is intricately related to STX1A expression. Furthermore, this experiment validates the previous use of1.0 μg hERG to 0.5 μg of STX1A cDNA / 30 mm dish for electrophysiology experiments as well as for subsequent molecular biology experiments (using HA-hERG) because of the moderate effect that STX1A imposes on HA-hERG at that dose. A weight ratio of 1:2 (hERG cDNA:

STX1A cDNA) for transfection represents a molar ratio of roughly 1 hERG channel for each 8 STX1A proteins, thereby ensuring the presence of excess STX1A (see calculation in Chapter 3: Materials and

Methods).

Pharmacological rescue of mutant or misfolded ion channels using high affinity pore blockers or pharmacological chaperone molecules has been well established as a means of increasing current density and cell surface expression (Sato et al., 1996; Gong et al., 2006). Naïve HEK 293 cells were transiently transfected with HA-hERG without or with STX1A and inhibition of hERG channel maturation by STX1A was assessed by detailed immunoblotting analysis. Fig. 22 A shows a typical

Western blot displaying two HA-hERG bands at 135 and 155 kDa. Lanes 3 and 4 represent cell lysates without and with STX1A cotransfection following 24-h incubation with 5 μM E-4031 supplemented in

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Figure 22. STX1A-mediated inhibition of HA-hERG channel maturation

A) Typical Western blot showing naïve HEK 293 cells transfected with HA-hERG without or with STX1A. Mature 155 kDa HA-hERG channel expression is markedly reduced when coexpressed with STX1A. Lanes 3 and 4 represent groups incubated with 5 μM E-4031 for 24 h. The channel-blocking drug partially restores mature HA- hERG protein expression. β-actin (Sigma, A5316, 1:20,000) was used as a loading control to demonstrate equal protein loading (lower panel). B) Densitometric analysis of mature HA-hERG protein expression / total HA-hERG protein. STX1A significantly reduces mature HA-hERG expression (n = 10 experiments, ** p < 0.01). E-4031 does not significantly rescue STX1A-mediated inhibition of HA-hERG channel maturation.

-87- the culture medium. Previously, I demonstrated that E-4031partially rescued alterations in whole-cell hERG current amplitude when coexpressed with STX1A (Fig. 18 A). In the present experiment, E-4031 caused a slight increase in mature HA-hERG protein relative to HA-hERG + STX1A alone. Fig. 22 B shows summarized densitometry data for the immunoblots. STX1A significantly reduced 155 kDa HA- hERG / total HA-hERG protein expression from 0.32 ± 0.03 to 0.16 ± 0.03 (p < 0.01, n = 10 experiments).

Incubation for 24 h with E-4031 partially restored mature HA-hERG protein following cotransfection with STX1A (HA-hERG + STX1A + E-4031 0.22 ± 0.04 vs. HA-hERG + E-4031 0.30 ± 0.04), however, this was not significantly different from HA-hERG + STX1A.

In addition to pharmacological rescue, a reduction in the temperature at which cells are incubated has also been shown to help stabilize protein folding and promote protein maturation and trafficking

(Zhou et al., 1999). For this reason, I also investigated the effect of incubating cells at reduced temperature. I hypothesized that reduced temperature might help stabilize folding and maturation, possibly impairing or disrupting the STX1A-mediated reduction in hERG channel maturation. Cells were transferred to a 30 °C incubator 24 h after transfection, and groups of cells transiently transfected with HA-hERG without or with STX1A cotransfection were incubated without or with 5 μM E-4031 for

24 h prior to harvesting for immunoblot analysis (Fig. 23 A). Remarkably, 24 h incubation of HA-hERG

+ STX1A cells alone is enough to restore normal mature HA-hERG levels (HA-hERG 0.38 ± 0.05 vs. HA- hERG + STX1A 0.31 ± 0.02, n = 4 experiments; not significantly different from HA-hERG; Fig. 23 B). This restoration of hERG protein maturation occurs independently of E-4031 and therefore is likely to rescue channel trafficking by an independent mechanism. Cells incubated with E-4031 were not statistically different from control groups (HA-hERG + E-4031 0.40 ± 0.03 vs. HA-hERG + STX1A + E-

4031 0.36 ± 0.02).

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Figure 23. Reduced temperature restores HA-hERG channel maturation

HEK 293 cells were transfected with HA-hERG without or with STX1A without or with 5 μM E-4031 incubation for 24 h. Incubation of samples at 30 °C instead of 37 °C facilitated the rescue of HA-hERG channel maturation. A) A typical immunoblot shows that mature HA-hERG channel expression still appears to be partially reduced by STX1A (lane 2), however, this effect is completely abolished by incubation for 24 h with E-4031. The lower panel shows the expression of α/β Tubulin (Cell Signaling 2148, 1:1000) producing a band at approximately 55 kDa, which was used as a loading control. B) Densitometric analysis of immunoblots demonstrate that there was no significant difference between the groups (n = 4 experiments).

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4.7 Truncated hERG proteins as tools for the characterization of hERG-STX1A interactions

Several hERG channel truncation mutations were obtained from the laboratories of Dr. Gail Robertson,

University of Wisconsin at Madison (hERG-Δ2-16, hERG-Δ2-354) and Dr. Alvin Shrier, McGill University

(HA-hERG-Δ814, HA-hERG-Δ860, HA-hERG-Δ860-899, HA-hERG-Δ899, HA-hERG-Δ960, HA-hERG-

Δ1000, HA-hERG-Δ1045, HA-hERG-Δ1120, HA-hERG WT). These truncation mutations are useful because several of them traffic to the plasma membrane and produce measurable hERG current. We hypothesized that these constructs could serve to not only validate some of the above experiments exploring the interaction of hERG and STX1A, but they may also help to identify the specific site of interaction between the residues. The hERG truncation mutations are schematically represented on a line diagram referring to a full-length hERG WT protein, which consists of 1159 amino acids (Fig. 24 A).

This schematic also illustrates residues bordering the cyclic nucleotide binding domain (cNBD) and

Per-Arnt-Sim (PAS) domains as well as 6 transmembrane segments (S1-S6). Arrows indicate the position of truncated hERG constructs. The C-terminal truncation mutations were numbered such that their number indicates the last residue removed from the truncated mutant channel. The truncations Δ2-16 and Δ2-354 are recognized by all anti-hERG antibodies directed toward the C- terminus, while all of the truncations provided by Dr. Shrier, including WT also contain an HA-tag in the distal N-terminus which allows them to be easily detected by anti-HA antibody. HEK 293 cells were transfected with equal amounts of cDNA for each truncation and Western blot analysis was performed (Fig. 24 B). Note that this Western blot was obtained using a combination of anti-HA and anti-hERG antibodies. Several truncations are retained by the ER-Golgi network as indicated by the absence of a mature protein band including hERG-Δ2-354, HA-hERG-Δ814, HA-hERG-Δ860, and HA- hERG-Δ860-899. Total hERG protein expression was examined qualitatively by (Fig. 24 C) revealing that several C-terminal truncations appear to express a larger quantity of protein than HA-hERG, however, it is inappropriate to compare the N-terminal truncations as they were exclusively detected with the anti-hERG antibody.

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Figure 24. Expression of hERG channel truncation mutations

A) Line diagram representing a full length hERG WT construct (1159 amino acid residues). Residues bordering the Per-Arnt-Sim (PAS) and cyclic nucleotide binding domain (cNBD) are indicated as well as the 6 transmembrane domains (S1-S6). Additionally, hERG truncation mutations appear below the line diagram with arrows indicating truncation locations. B) Western blot was used to demonstrate the relative size and expression levels of the various hERG truncation mutations compared with HA-hERG WT controls (lanes 1 and 12). Two N- terminal truncation mutations were recognized solely by an anti-hERG antibody directed against the C-terminus whereas all of the other constructs contained an HA-tag which was detected with an anti-HA antibody. Constructs with double bands represent channels that are capable of trafficking to the plasma membrane. C) Densitometric analysis of the above Western blot bands reveals unequal protein expression among the truncation mutations. HA-hERG-Δ1120 appears to have the strongest expression level of all the constructs. (Note that it is not appropriate to compare the two N-terminal truncation mutations because they are only detected by the anti-hERG antibody and do not possess HA-tags).

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The C-terminal truncation mutation HA-hERG-Δ1120 qualitatively appears to express a larger ratio of mature complex-glycosylated hERG protein relative to control HA-hERG (see Fig. 24 B). Therefore, I hypothesized that this would be a good model to compare with the results obtained in earlier experiments demonstrating the effects of STX1A-mediated reduction in hERG channel maturation.

Western blot analysis was used to study the dose-dependent reduction of mature HA-hERG-Δ1120 protein caused by cotransfection with STX1A (Fig. 25 A). The ratio of mature/ total protein expression for HA-hERG-Δ1120 was reduced from 0.44 from control (no STX1A cotransfection) to 0.32 with 0.5 μg

STX1A and further to 0.04 with 5.0 μg STX1A cotransfection (Fig. 25 B). STX1A expression was immunoblotted for each lysate sample (Fig. 25 A, lower panel), quantified using densitometry and plotted versus STX1A transfection amount in arbitrary densitometric values on the right y-axis. STX1A expression could be detected for each transfection level, with lower concentrations requiring prolonged exposure of the chemiluminescent signal. This result demonstrates that the truncation mutant construct HA-hERG-Δ1120 yields a larger proportion of mature hERG protein to total protein, which is reduced with increasing amounts of STX1A cotransfection comparable to HA-hERG WT and

STX1A.

The C-terminal truncations HA-hERG-Δ1120, HA-hERG-Δ1045, HA-hERG-Δ1000, HA-hERG-Δ960 and

HA-hERG-Δ899 are capable of trafficking to the plasma membrane. These constructs were transfected with and without STX1A for immunoblot analysis to determine what effect STX1A would have on their complex-glycosylated mature hERG expression (Fig. 26). We hypothesized that if STX1A functionally interacts with a region of hERG upstream of the C-terminus, we would observe no rescue of the hERG maturation deficiency caused by STX1A. If STX1A, however, interacts with a region of the hERG C- terminus upstream of residue 899, we should be able to observe the functional inhibition of the

STX1A-mediated hERG trafficking deficiency. When coexpressed with STX1A, the majority of the truncation mutants tested demonstrated a reduction in mature hERG protein (Fig. 26 A). An exception is HA-hERG-Δ899 in which the mature band intensifies and shifts to a higher molecular weight. This requires further study, but could be the result of HA-hERG-Δ899 channels strongly interacting with

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Figure 25. STX1A inhibits HA-hERG-Δ1120 maturation in a dose-dependent manner

HA-hERG-Δ1120 was used to verify the dose-dependent impairment of hERG-channel maturation because of its proportionately larger expression of mature HA-hERG-Δ1120 protein to total HA-hERG-Δ1120 protein. A) Western blot shows HA-hERG-Δ1120 channel expression without or with increasing cotransfection with STX1A (top panel). The upper (mature) HA-hERG-Δ1120 band is reduced with increasing STX1A coexpression (lower panel). The last lane demonstrates that naïve HEK 293 cells do not possess endogenous HA-hERG-Δ1120 or STX1A protein. B) Densitometric analysis of the above Western blot. HA-hERG-Δ1120 expression is plotted as a ratio of mature to total HA-hERG-Δ1120 protein relative to STX1A transfection dose. STX1A expression is plotted on the right y-axis as arbitrary densitometric units, which increase with larger transfection doses.

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Figure 26. hERG and STX1A functionally interact downstream of residue 1000 hERG truncation mutations were used to assess STX1A-mediated inhibition of hERG channel maturation. A) Typical Western blots depicting HA-hERG WT or HA-hERG truncation mutations without (-) or with (+) STX1A cotransfection. STX1A reduced mature hERG expression in HA-hERG WT, HA-hERG-Δ1120, HA-hERG-Δ1045 and HA-hERG-Δ1000 constructs, and had no effect on HA-hERG-Δ960. Mature HA-hERG-Δ899 protein expression actually increased as a result of STX1A coexpression. B) Summarized densitometric analysis of n = 4-10 experiments per group representing mean data for the ratio of mature hERG to total hERG expression, without or with STX1A coexpression. (** p < 0.01; * p < 0.05).

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STX1A protein despite the harsh SDS detergent conditions. Multiple densitometric analyses of several

Western blots for each experimental group were pooled and are depicted in Fig. 26 B. Mature hERG protein is expressed as a ratio of the mature hERG band to the total hERG protein expression. HA- hERG expression was significantly reduced by STX1A from 0.35 ± 0.03 to 0.17 ± 0.03 (p < 0.01, n = 10 experiments). HA-hERG-Δ1120 expression was significantly reduced from 0.32 ± 0.01 to 0.18 ± 0.04 (p

< 0.05, n = 4 experiments). HA-hERG-Δ1045 was significantly reduced from 0.32 ± 0.03 to 0.17 ± 0.03

(p < 0.05, n = 4 experiments) and HA-hERG-Δ1000 was reduced by STX1A cotransfection from 0.41 ±

0.01 to 0.25 ± 0.02 (p < 0.05, n = 4 experiments). Interestingly, while there was not a significant reduction in HA-hERG-Δ960 (0.38 ± 0.05 vs. 0.30 ± 0.02 (STX1A); n = 4 experiments), surprisingly as mentioned above, hERG-HA-Δ899 actually increased the proportion of mature hERG protein from 0.40

± 0.02 to 0.60 ± 0.01 when coexpressed with STX1A (p < 0.01, n = 4 experiments). These results suggest that hERG and STX1A functionally interact upstream of residue 1000.

Previously, we observed that E-4031 appears to qualitatively increase mature HA-hERG levels when coexpressed with STX1A, however, the effect is not dramatic enough to be statistically significant. We hypothesized that use of the more distal C-terminal truncation mutations may serve as an alternative diagnostic tool for studying the inhibitory effects of STX1A on mature hERG expression. We have shown that the constructs HA-hERG-Δ1120 and HA-hERG-Δ1045 functionally interact with STX1A resulting in significantly reduced mature hERG expression (Fig. 26). Western blot analysis was performed on lysates obtained for groups of HEK 293 cells transfected with either of the C-terminal truncations without or with STX1A. Western blots were also performed for samples treated with E-

4031 for 24 h prior to protein isolation. Fig. 27 A and C show typical Western blots obtained for HA- hERG-Δ1120 and HA-hERG-Δ1045, respectively. STX1A reduces the expression of mature HA-hERG protein in both truncation mutants, while incubation with E-4031 appears to qualitatively restore control levels in only the HA-HERG-Δ1045 construct. Fig. 27 B shows summarized densitometric analysis of 3 experiments for HA-hERG-Δ1120. The ratio of mature HA-hERG-Δ1120 protein to total

HA-hERG-Δ1120 protein was not significantly increased by 24 h E-4031 incubation (HA-hERG-Δ1120 +

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Figure 27. E-4031 rescues STX1A-dependent inhibition of hERG-HA-Δ1045 maturation

The C-terminal truncation mutations hERG-HA-Δ1120 and hERG-HA-Δ1045 were used to study the effect of STX1A cotransfection and the previously observed rescue mechanism of E-4031. A) Typical Western blot showing hERG-HA-Δ1120 expression without (lane 1) and with (lane 2) STX1A expression, and 24 h E-4031 expression. B) Densitometric analysis of 3 experiments reveals that STX1A-mediated reduction of mature hERG protein is not significantly rescued by 24 h E-4031 incubation. C) Typical Western blot for hERG-HA-Δ1045 with the same conditions described for A). D) Averaged densitometric analysis of n = 3 experiments. E-4031 incubation significantly restores the ratio of mature hERG to total hERG protein. (*p < 0.05)

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STX1A 0.19 ± 0.07 vs. HA-hERG-Δ1120 + STX1A + E-4031 0.27 ± 0.05). However, Fig. 27 D illustrates that mature HA-hERG-∆1045 protein expression was significantly increased following 24 h E-4031 incubation (HA-hERG-∆1045 + STX1A 0.12 ± 0.02 vs. HA-hERG-∆1045 + STX1A + E-4031 0.37 ± 0.04, n =

3 experiments, p < 0.05). Moreover, HA-hERG-∆1045 + STX1A + E-4031 was not significantly different from HA-hERG-∆1045 + E-4031 in the absence of STX1A (0.45 ± 0.05). Cumulatively, we have shown that STX1A-mediated inhibition of mature HA-hERG is partially restored by 24 h incubation with E-

4031, with the most dramatic example being the HA-hERG-Δ1045 construct. These results support our observation that STX1A-induced current amplitude reductions can be partially rescued following incubation with the hERG channel blocker E-4031 (see Fig. 18).

4.8 hERG and STX1A binding experiments

STX1A binding with hERG was investigated by utilizing a GST pull-down assay with recombinant protein. hERG was expressed in tsA 201 cells and cell lysate from transfected cells was incubated with

GST alone or with GST-STX1A bound to glutathione-agarose beads. hERG bound GST-STX1A but not

GST demonstrating that STX1A can directly bind to hERG in our cell system (Fig. 28 A). Next, we also examined which domain of STX1A specifically interacts with hERG. The STX1A active domain has been

2+ shown to functionally interact with N-type Ca channels and KV2.1 channels (Wiser et al., 1996; Leung et al., 2005). Normally, the protein exists in a ‘closed’ configuration in which the N-terminal HABC domain folds over to inhibit the C-terminal H3 domain from assembling with the SNARE machinery.

Therefore, we performed pulldown experiments with truncated versions of STX1A comprising its HABC domain (corresponding to amino acids 1-160) and H3 domain (amino acids 191-256). As expected, hERG bound preferentially to the STX1A-H3 domain, which is consistent with the idea that when it becomes activated, it adopts an ‘open’ configuration, allowing this domain to become available for the interaction with hERG. hERG cell lysate was used for control and the primary antibody was directed against the C-terminus.

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Figure 28. Interaction of hERG and STX1A

Two types of experiments were carried out to test hERG and STX1A interaction. A) GST-pulldown assay utilizing recombinant protein investigated STX1A binding with hERG. hERG was expressed in tsA 201 cells and cell lysate was incubated with GST alone or with GST-STX1A bound to glutathione-agarose beads. hERG bound GST-STX1A but not GST demonstrating that STX1A can bind directly to hERG. Pulldown experiments with GST fusion proteins of the STX-HABC and STX-H3 domains demonstrated that the H3 domain preferentially bound to hERG. Immunoblot analysis of hERG protein was with an anti-hERG antibody directed against the C-terminus (Alomone, APC-062, 1:2000). B) Coimmunoprecipitation of hERG with STX1A was examined by coexpression of the proteins in HEK 293 cells. Lysates were incubated with STX1A antibody (Sigma S0664, 2 μL) and Protein-A- Sepharose beads. Samples were eluted by boiling and immunoblotted with anti-hERG primary antibody (Alomone, APC-062, 1:2000), revealing the expression of one band at 135 kDa. Negative control (lane 2) contained no STX1A antibody during binding, and positive control was regular hERG + STX1A cell lysate.

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GST-pulldown experiments are excellent for establishing whether two proteins can physically bind, but due to the nature of “protein baiting”, this assay does not test whether our proteins of interest are actually interacting in our mammalian cell system. Therefore, we sought to test whether hERG and

STX1A physically interact using coimmunoprecipitation. HEK 293 cells were cotransfected with both hERG and STX1A cDNA constructs and grown for 48 h prior to harvesting. Samples were precleared with Protein-A-Sepharose beads to eliminate any non-specific binding. Samples were then incubated with anti-STX1A antibody and beads, and then washed thoroughly. Boiling the samples eluted protein and Western blot was subsequently performed. Fig. 28 B shows that the STX1A immunoprecipitation sample immunoblotted for hERG protein reveals the expression of one band at

135 kDa. Lysate was saved prior to binding for a positive control. Another sample was prepared containing no STX1A antibody, which was used as a negative control (lane 2). Overall, this result illustrates the interaction of hERG and STX1A in our cell system.

Coimmunoprecipitation of STX1A and truncated hERG proteins was used to assess a possible site of interaction (Fig. 29 and 30). HEK 293 cells cotransfected with STX1A and hERG constructs were coimmunoprecipitated with anti-STX1A or anti-HA and subsequently immunoblotted with anti-HA, anti-hERG or anti-STX1A. STX1A interacts strongly with the N-terminal truncations hERG-Δ2-16 and hERG-Δ2-354 (Fig. 29 A) and the C-terminal truncations HA-hERG-Δ814 and HA-hERG-Δ860 (Fig. 29 B).

These results show that the site of interaction between hERG and STX1A must occur between the amino acid residues 354 and 814, possible with an intracellular loop or transmembrane region.

However, this may not be the only site of interaction as our functional data suggest that hERG and

STX1A may interact upstream of residue 1000 (Fig. 26). Next, the relative binding strengths of the C- terminal truncations were assessed. The presence of a common N-terminal HA-tag makes this possible. Fig. 30 A illustrates all of the cell lysate samples for the two sets of experiments. The top panel shows that following pre-clearing with beads, HA-hERG expression is not consistent between truncation groups despite consistent α/β tubulin loading controls (second panel). Also note that with prolonged exposure there is a prominent unspecific band at 125 kDa in HEK cells. STX1A expression in

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Figure 29. hERG and STX1A interaction occurs between the residues 354 and 814

Coimmunoprecipitation of hERG truncation mutations was used to identify the site of interaction between hERG and STX1A. A) The two N-terminal hERG truncation mutations hERG-Δ2-16 and hERG-Δ2-354 were immunoprecipitated with anti-STX1A antibody (Sigma S0664, 2 μL) and immunoblotted with anti-hERG primary antibody (ABR PA3-860, 1:3000). Both truncation mutations strongly bind to STX1A. Positive controls were cell lysates, which were set aside prior to immunoprecipitation. B) The two largest C-terminal truncation mutations were immunoprecipitated with anti-STX1A antibody (Sigma S0664, 2 μL) and then immunoblotted with anti-HA (Sigma H3663, 1:5000) to test HA-hERG truncation expression. Both truncation mutants coimmunoprecipitated with hERG. (ф) represents negative controls omitting STX1A primary antibody during immunoprecipitation, and positive controls were whole cell lysates set aside prior to immunoprecipitation.

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Figure 30. STX1A and hERG interaction is strongest with the shortest C-terminal truncations

The relative strength of the C-terminal HA-hERG truncation mutation interactions with STX1A was assessed using coimmunoprecipitation. A) Western blots show lysate controls for both sets of experiments. Top panel shows HA-hERG expression (which is inconsistent between groups), second panel shows α/β tubulin loading controls for HA-hERG lysates, third panel shows STX1A lysate controls (which also show inconsistent expression between groups after pre-clearing with beads), and the bottom panel shows the α/β tubulin loading controls for STX1A lysates. B) Coimmunoprecipitation results illustrate that the shortest C-terminal truncation mutations interact most strongly with STX1A. HA-hERG-Δ814 and HA-hERG-Δ860 interactions appear to be weakest of all, but may be due to their extremely low expression levels. IP: Φ indicates negative controls omitting the use of primary antibodies during immunoprecipitation (second and forth panels).

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Coimmunoprecipitation demonstrates that the strongest interactions occur between larger C-terminal truncations and STX1A, while HA-hERG-Δ814 and HA-hERG-Δ860 interactions are much weaker, likely due to their decreased expression levels. Negative controls are shown in the second and forth panels in which immunoprecipitation samples did not include primary antibodies. Overall, these results suggest that the major site of hERG-STX1A interaction lies between residues 354 and 814.

4.9 Endogenous expression of hERG & STX1A

In order to assess whether the interaction between hERG and STX1A exerts a physiologically relevant role, the endogenous expression of those proteins was examined. HL-1 heart cells (a mouse atrial cell line) have been shown to possess a rapidly activating delayed rectifier current (Akhavan et al., 2003). Confluent dishes of HL-1 cells were harvested in 0.5% NP-40 lysis buffer and Western blots were performed as described above. Fig. 31 A shows that HL-1 cells express both immature and mature forms of hERG. In this immunoblot, hERG-HEK 293 cells were used as control. hERG expression was also examined in mouse and rat heart tissue samples. The methods employed for isolation of endogenous cardiac protein are outlined in the materials and methods section (Chapter 3). Fig. 31 B illustrates a Western blot examining the expression of hERG protein in rat brain (lanes 1,2), hERG-HEK

293 cells (lane 3), HL-1 cells (lane 4), mouse heart membrane lysate (lanes 5,6) and whole rat heart lysates (lanes 7,8). Signals were detected with the erg1 antibody, which consists of a 15 amino acid epitope which has identical homology to mouse and is directed against the C-terminus. We detected the expression of hERG protein in rat brain, HL-1 cells, hERG-HEK-292 cells (control) and rat heart, but not mouse heart membrane. Following prolonged exposure, double bands were detected for hERG-

HEK 293 cells and HL-1 cells, but results for the other tissue types were unclear. We also detected the

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Figure 31. Endogenous expression of hERG and STX1A

HL-1 heart cells (a mouse atrial line), mouse heart membrane, and whole rat heart was used to assess the endogenous expression of hERG (A, B) and STX1A (C). (A) hERG expression is visualized as a double band at approximately 135 and 155 kDa in HL-1 cells using an anti-hERG antibody directed against the C-terminus (Alomone APC-016, 1:250). hERG-HEK 293 cells served as a positive control. B) hERG expression is also detected at 135 kDa in rat brain and whole rat heart lysate using the erg1 antibody (Alomone APC-016, 1:400). hERG was not detected in mouse heart membrane using any of the antibodies tested. C) STX1A expression has been previously shown in rat heart and rat brain lysates. Here, rat brain serves as a positive control. STX1A produces a doublet of bands that represent STX1A and STX1B, which cannot be distinguished by the STX1 monoclonal antibody (Sigma S0664, 1:1000) used in this experiment. Mouse heart membrane and HL-1 cells both express STX1A.

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expression of hERG protein in the same cell types with another antibody directed against the C- terminus. While hERG expression in whole rat heart lysate is very strong, expression in mouse heart membrane is absent. This led us to question the quality our membrane preparation, which should have concentrated hERG protein expression. We tested our preparation and verified its purity with antibodies directed against the Na+/K+ ATPase, which is membrane specific, and GAPDH, a cytosolic protein marker. We were also unable to detect hERG expression in mouse heart membrane using two other commercially available antibodies (Santa Cruz C-20 and N-20).

In addition to investigating the endogenous expression of hERG protein, the endogenous expression of STX1A was investigated in HL-1 cells, mouse and rat heart. Fig. 31 C illustrates the detection of

STX1A expression in lysates of mouse heart membrane, rat brain (positive control) and HL-1 cells using a monoclonal antibody. This antibody reveals the expression of a doublet representing STX1A and

STX1B isoforms (Sigma anti-STX1A antibody is not isoform specific). Comparable results were previously reported demonstrating the expression of both STX1 isoforms in rat brain, whole rat heart and rat heart membrane preparations (Kang et al., 2004). Furthermore, STX1 expression was identified in whole intact cardiomyocytes using confocal microscopy. Overall, these results suggest that STX1A is present in both primary mouse and rat heart cells and the HL-1 cell line. Furthermore, these results suggest that investing more time into the study of an endogenous hERG and STX1A interaction is warranted in case of HL-1 cells and isolated rat heart lysate. Furthermore, HL-1 cells may provide a better model for endogenous protein interaction because of their ease of use, abundant availability, and high hERG expression levels.

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Chapter 5: Discussion

The purpose of this study was to determine the extent to which the SNARE protein syntaxin 1A

(STX1A) interacts with the cardiac voltage-gated K+ channel hERG. STX1A has been shown to regulate the PM trafficking of a number of ion channels including CaV and KV channels in both secretory and nonsecretory cells (Bezprozvanny et al., 2000; Leung et al., 2007). We decided to pursue this study because of observations involving the interaction of STX1A with a variety of K+ channels, including the cardiac KATP, KV4.2 and KV4.3 channels (Kang et al., 2004; Ahmed et al., 2007; Yamakawa et al., 2007). The expression and activity KV4.2 and KV4.3 channels, which are highly expressed in the heart, are known to be regulated by numerous cytosolic and membrane associated auxiliary subunits, making these channels ideal candidates for interaction with STX1A (Yamakawa et al., 2007). hERG channels do not appear to interact as promiscuously with ancillary or β-subunits; however, they have been shown to be regulated by numerous chaperone protein molecules and are hypothesized to interact with the

MiRP1 β-subunit in vivo, although this remains controversial (Abbott et al., 2007).

In order to assess the potential protein-protein interaction between hERG and STX1A, I utilized a multi-disciplinary approach involving the use of electrophysiological and molecular biochemical assays. Experiments in this study were designed around five main aims listed in Chapter 2. To summarize, I have assessed the effect of STX1A on hERG channel gating and kinetics, channel trafficking and cell surface localization, as well as the binding sites of the two proteins using full length and truncated protein constructs. Additionally, I have shown that STX1A-mediated hERG channel inhibition can be disrupted using hERG channel blockers and reduced temperature. Finally, I have investigated the endogenous expression of these proteins in order to better establish a physiologically relevant link regarding their interaction. These results have important implications for defining the intrinsic biological mechanism by which SNARE proteins regulate the expression and function of K+ channels, particularly in the heart.

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5.1 hERG channel expression in HEK 293 cells is an appropriate model system

In the present study, I have selected HEK 293 cells, derived from human embryonic kidney cells to study the interaction of the proteins STX1A and hERG. These cells were selected over tsA-201 cells because of their low levels of endogenous outward current (see Fig. 11 B and Fig. 12 C) (Zhou et al.,

1998b). The initial detailed hERG channel characterization was achieved by utilizing Xenopus oocytes or transient expression of hERG channels in HEK 293 cells (Sanguinetti et al., 1995; Trudeau et al., 1995;

Snyders and Chaudhary, 1996). Careful electrophysiological assessment of hERG channel gating and kinetics revealed that hERG encodes a K+ channel which closely resembles but does not fully recapitulate the properties of the native IKr in the heart (see Chapter 1.3.3) (Sanguinetti and Tristani-

Firouzi, 2006). For example, native IKr is characterized by approximately 10x faster activation and deactivation kinetics than hERG channels expressed in mammalian cells (Sanguinetti, 1999). Current hypotheses assert that these differences in hERG channel function can be resolved by interaction of native hERG channels with the MiRP1 β-subunit, or by heteromeric assembly of WT hERG channels with the N-terminal truncation splice variant HERG-1b (Abbott et al., 1999; Jones et al., 2004; Sale et al.,

2008). Despite this, use of stably transfected hERG-HEK 293 cells has become the most widely used and accepted method of testing the biochemical, electrophysiological and pharmacological properties of hERG channels (Zhou et al., 1998b; Mitcheson, 2008). This is particularly relevant in light of the S7B guideline (http://www.ich.org) issued by the International Conference on Harmonization of

Technical Requirements for Registration of Pharmaceuticals for Human Use. This guideline recommends the in vitro evaluation of new experimental compounds for unintentional inhibition of hERG channel function (Zeng et al., 2008). High-throughput screening and automated planar patch- clamp technologies used for hERG channel screening also utilize stably-transfected mammalian cell lines including HEK 293 cells (Netzer et al., 2003; Guo, L. and Guthrie, 2005). Overall, in light of our current, albeit limited understanding of native IKr, expression of hERG channels in mammalian cells is the most reproducible and closest approximation for testing the physiological properties of these channels.

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The use of stably transfected cell lines assures high hERG protein expression levels, relatively low background currents and a high degree of uniformity of current amplitude from cell to cell. HEK 293 cells are also a particularly appealing cell line to utilize because of their fast reproduction and easy maintenance, high transfection efficiency, and relatively small cell size allowing for easy and reliable electrophysiological measurements (Thomas and Smart, 2005). We obtained stably transfected hERG-

HEK 293 cells from the laboratory of Dr. Craig January at the University of Wisconsin-Madison in early

2006, however, preliminary electrophysiological assessment of the hERG-STX1A interaction was carried out using transiently transfected tsA-201 cells which possess significant background endogenous currents (Appendix 1) (Varghese et al., 2006). hERG channel current amplitude was found to be optimal following transfection of 1.0 μg of cDNA per 30 mm dish. STX1A cotransfection was 0.5

μg of cDNA, therefore ensuring a molar ratio of 1 hERG:8 STX1A functional proteins (see Chapter 3.4).

Although we could not predict the stoichiometry of the hERG-STX1A interaction, we believe that providing excess STX1A protein optimized the likelihood of a detectable protein-protein interaction.

Upon switching to use of stably transfected hERG-HEK 293 cells for electrophysiological assessment, we continued to use 0.5 μg of STX1A cDNA per 30 mm dish. This decision was supported by the fact that whole-cell hERG current amplitude was actually smaller in the stable cell line versus transient transfection, implying that less hERG protein is available at the plasma membrane and thereby allowing for an even greater molar ratio of hERG to STX1A protein available for interaction.

5.2 STX1A-dependent reduction of hERG channel open probability

Previous reports exploring the relationship between SNARE proteins and K+ channels have revealed that regulation of K+ channel function is highly specific depending on the SNARE protein and channel in question. It is difficult to make universal conclusions regarding the mechanism by which STX1A regulates the functionality of K+ channels; however, it appears as if STX1A expression generally reduces K+ channel function, resulting in a reduction of whole-cell current amplitude. Whole-cell

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hERG current (IhERG) is the product of three factors: 1) the number of functional hERG channels at the

PM (n), 2) the probability that an individual channel is open at a given time (PO); and 3) the amplitude of single-channel conductances (i). Measurement of single-channel conductances requires careful single-channel recordings which are technically complex and beyond the scope of this project. One can, however, assess the gating properties of a channel by performing whole-cell patch clamp experimentation. Use of carefully designed voltage-protocols can assist in the determination of hERG channel kinetics and gating properties, which can help define channel open probability. Additionally, whole-cell patch clamping provides a direct measurement of hERG channel current amplitude, which is reflective of the number of functional channels at the PM.

The first aim of this project was to determine the extent to which STX1A regulated the function of hERG channels by utilizing electrophysiological measurements of whole-cell hERG-HEK 293 cell currents. STX1A significantly reduced peak hERG current amplitude and tail currents without affecting the C-type inactivation or rectification properties of the channel. Surprisingly, despite the large reduction in hERG current amplitude, no change in the time constants of channel activation, deactivation, fast inactivation or recovery from inactivation was observed. These results imply that the

STX1A-mediated reduction of whole-cell hERG current amplitude is likely not due to a significant change in hERG channel open probability. This contrasts with observations from previous studies

+ including at least 5 KV channels, in which STX1A-mediated impairment of K channels was at least partially due to changes in channel gating and kinetics.

Table V summarizes data obtained from detailed analysis of the STX1A-dependent modulation of KATP,

KV1.1, KV1.2, KV2.1, KV4.2 and KV4.3. STX1A reduces current amplitude in each of these channels, albeit via distinct mechanisms which cannot be easily generalized. For example, STX1A decreases activation kinetics in KV1.2 and KV2.1 (Tsuk et al., 2005; Neshatian et al., 2007), while it increases the rates of fast inactivation and deactivation in KV1.1 and KV4.2, respectively (Fili et al., 2001; Yamakawa et al., 2007).

These changes in channel gating kinetics act to reduce the open probability of the aforementioned K+

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Table V: Summary of syntaxin 1A interaction with K+ channels

Channel Current Steady-State Other Kinetics Binding Reference Amplitude Inactivation KATP ↓ N/A ↓ activation SUR1 & (Cui, N. et al., 2004; SUR2A Kang et al., 2004; Pasyk subunits et al., 2004) KV1.1 ↓ N/A ↑ fast inactivation N-term (Fili et al., 2001; Ji et al., 2002b)

KV1.2 ↓ N/A ↓ activation; N-term (Ji et al., 2002a;

→ shift in V1/2 of Neshatian et al., 2007) activation KV2.1 ↓ ↓ slope factor; ↓ activation; C-term (Leung et al., 2003;

← shift in V1/2 ← shift in V1/2 of Michaelevski et al., (-20.1 mV) activation 2003; Tsuk et al., 2005)

KV4.2 ↓ → shift in V1/2 ↑ deactivation; N-term (Yamakawa et al., 2007) (5.2 mV) ↑ recovery from inactivation

KV4.3 ↓ ← shift in V1/2 none ? (Ahmed et al., 2007) (-3.1 mV)

hERG ↓ ← shift in V1/2 none C-term (Mihic et al., 2006; (-13.3 mV) Mihic et al., 2008)

← left shift (hyperpolarizing); → right shift (depolarizing); ↑ increases (or accelerates kinetics); ↓ decreases (or slows kinetics)

-109- channels, resulting in significantly reduced whole-cell current amplitude. Structural differences in the cytoplasmic N- and C-terminal domains likely account for the unique patterns of STX1A-mediated modulation of K+ channel currents, and so without a detailed understanding of K+ channel-STX1A binding structure, it is difficult to develop a model explaining how interaction of the proteins is translated into the specific regulation of channel gating kinetics.

hERG channel inactivation is caused by a C-type inactivation mechanism (see Chapter 1.3.2) which is voltage-dependent and stronger at more depolarized potentials giving the channels its inward rectification property (Smith et al., 1996; Wang, S. et al., 1997). Despite an apparent inability for STX1A to affect the kinetics of hERG channel gating, it did induce a hyperpolarizing shift in the midpoint voltage of steady-state inactivation. This parameter reflects the increasingly reduced availability of hERG channels at progressively more depolarized potentials, resulting in enhanced inward rectification of the channel. Following cotransfection with STX1A, hERG-HEK 293 cells undergo a subtle but significant -13.3 mV hyperpolarizing shift in the V1/2 of steady-state inactivation. More simply, this means at a given test potential, control hERG cells will have more channels available to open than hERG + STX1A cells. This can partially explain the mechanism by which STX1A reduces whole-cell hERG currents, although we hypothesized that another more predominant mechanism may underlie this dramatic reduction in whole current amplitude.

STX1A also causes significant shifts in steady-state inactivation in KV2.1, KV4.2 and KV4.3 (see Table V).

Similar to hERG channels, STX1A causes a large hyperpolarizing shift (~20 mV) in the midpoint of KV2.1 steady-state inactivation (Michaelevski et al., 2003; Tsuk et al., 2008). Electrophysiological measurements of KV2.1 currents were taken from Xenopus oocytes, and steady-state inactivation was determined following 25- and 5-s depolarizing prepulses at incremental test potentials completely inactivating channels, and then subsequently depolarizing to +50 mV for 120 ms for construction of the inactivation curve (Michaelevski et al., 2003). In contrast to the large shift in KV2.1 steady-state inactivation, KV4.2 and KV4.3 display very slight alterations in this parameter (Ahmed et al., 2007;

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Yamakawa et al., 2007). Additionally, despite their structural similarities, these channels appear to be differentially modulated by STX1A such that KV4.2 undergoes a slight depolarizing shift and KV4.3 undergoes a slight hyperpolarizing shift in the midpoint of steady-state inactivation. Despite their structural similarities, STX1A regulates channel function differently in these two KV4.x family members, and it is unlikely that such subtle alterations in steady-state inactivation have significant functional implications for whole-cell channel currents. Therefore, I believe that the mechanism governing the

STX1A-mediated hyperpolarizing shift in the steady-state inactivation of hERG channels may resemble that of KV2.1 channel modulation. In order to establish this, a more detailed understanding of the protein-protein binding structure is required.

In 1998 and 2002, two Japanese laboratories identified novel missense mutations in hERG which resulted in familial long QT-syndrome caused by a reduction in hERG current amplitude and alterations in the voltage dependence of channel inactivation (Nakajima et al., 1998; Hayashi, K. et al.,

2002). All three of these mutant hERG channels, V630L, A614V and E637K were unable to form functional channels when expressed alone, but were capable of trafficking to the PM and produced measurable current following heterologous expression with WT hERG. Remarkably, all of these missense mutations resulted in a large reduction in hERG current amplitude, as well as producing a hyperpolarizing shift in the midpoint of steady-state inactivation. The A614V mutation caused a mild -

9.3 mV shift in the midpoint of hERG channel steady-state inactivation while having no affect on the kinetics of channel activation, deactivation, inactivation or recovery (Nakajima et al., 1998). V637K resulted in a more severe hyperpolarizing shift in steady-state inactivation (-22.2 mV) as well as accelerating the rate of fast inactivation and recovery from inactivation (Nakajima et al., 1998). E637K resulted in a -16.4 mV hyperpolarizing shift in steady-state inactivation as well as an +8.8 mV depolarizing shift in the midpoint of steady-state activation without affecting other gating kinetic parameters (Hayashi, K. et al., 2002). Taken together, these results clearly demonstrate that the voltage dependence of hERG channel inactivation can be altered or modulated independently from the voltage dependence of channel activation, as well as from alterations in gating kinetics. These

-111- findings represent a novel mechanism for reduction of hERG channel current causing LQT2 by altering the channel’s inactivation voltage sensitivity. A reduction in whole-cell current by this mechanism is an important determinant of phase 3 cardiac AP duration (Nakajima et al., 1998; Zhou et al., 1998b).

These findings also provide information about the molecular basis of hERG channel kinetics.

Mutations affecting the channel pore have demonstrated that the residues A614-E637 may be related to conformational changes during hERG channel inactivation (Smith et al., 1996; Hayashi, K. et al.,

2002). In particular, the residues G628-S631 are believed to be part of the hERG channel inactivation gate with residue V630 being a particularly important residue for the voltage dependence of hERG channel inactivation (Schonherr and Heinemann, 1996; Smith et al., 1996). To support this, the missense mutation S631C causes acceleration of fast inactivation kinetics, and S631A results in slowing of fast inactivation kinetics (Zou et al., 1998).

Careful consideration of the above evidence has led us to hypothesize that inhibition of hERG channel currents by STX1A is at least partially due to the inhibitory effect of hyperpolarization of steady-state inactivation. Modulation of this parameter may be analogous to the mechanism by which STX1A impairs KV2.1 currents (Michaelevski et al., 2003), which is further bolstered by the finding that STX1A interacts with the C-terminus of KV2.1 (Leung et al., 2003). This is in agreement with the finding that specific residues located in the channel pore and S6 helices have important implications for voltage- sensitivity of hERG channel inactivation and C-type inactivation in general. Therefore, we further hypothesized that STX1A and hERG likely interact near the cytosolic C-terminal domain causing this secondary mechanism for inhibition of channel function by directly or indirectly interacting with the

S6 domains. Additionally, we hypothesized that the primary mechanism by which STX1A impairs hERG channel function is by reducing the number of functional channels present at the PM.

5.3 STX1A-mediated reduction in PM expression of hERG channels

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Following exhaustive electrophysiological analysis of the hERG-STX1A interaction, I shifted the focus of this research program to characterizing this protein-protein interaction by means of molecular biochemistry. The second aim of this research project was to evaluate STX1A-induced changes in hERG channel trafficking. Immunocytochemistry and confocal microscopy revealed that control hERG-HEK 293 cells display extensive cytoplasmic expression of hERG protein as well as distinct but less predominant PM expression. Following STX1A coexpression, the PM expression of hERG protein was markedly reduced while both proteins were strongly colocalized with periplasmic distribution within highly dense intracellular compartments. Therefore it is likely that STX1A and hERG directly interact resulting in the trafficking-deficient phenotype producing the significant decrease in whole- cell hERG currents discussed above. Moreover, coimmunofluorescence demonstrates that STX1A appears to trap hERG channels in highly-dense organelles in the periplasmic space. It is surprising that

STX1A expression appears to be primarily confined to the cytoplasmic space, especially considering its most common role as a target-membrane SNARE protein (Jahn and Scheller, 2006). Furthermore, our lab and our collaborator, Herbert Gaisano, have identified the PM expression of STX1A in both rat

(Kang et al., 2004) and mouse (Ng et al., 2008) cardiomyocytes, with weaker cytoplasmic expression. It is possible that in our overexpression system, STX1A is capable of functioning at both “donor” and

“acceptor” membranes, moving in both anterograde and retrograde directions (Sollner et al., 1993). It is interesting to note that the PM distribution of STX1A when coexpressed with KV2.1 is high relative to the cytoplasm, however, STX1A also serves to significantly reduce the PM expression of this K+ channel

(Leung et al., 2003; Leung et al., 2005).

Several other groups have successfully utilized confocal microscopy for the qualitative and quantitative assessment of hERG channel PM expression. One of the first such studies investigated the trafficking-deficient LQT2 mutant T65P (Paulussen et al., 2002). Cleverly, this study used two fluorescent tags, one to assess expression of the T65P hERG mutant, and another, a red fluorescent ER vector, to define the ER expression pattern of the LQT2 mutant channels. Using HEK 293 cells, this study clearly demonstrated the ER-retention of the trafficking-deficient protein. Following this, the

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G601S (Delisle et al., 2003) and the G604S (Huo et al., 2008) LQT2 mutants were also shown to produce trafficking-deficient phenotypes resulting in ER retention and accumulation. The latter group fluorescently tagged calnexin, an ER marker protein, clearly demonstrating the coimmunofluorescence of these proteins (Huo et al., 2008).

Based on similarities in the periplasmic distribution of hERG and STX1A with these LQT2 mutant expression patterns, we hypothesize that the intracellular compartment in which trafficking-impaired hERG channels reside is in fact the ER. Further experiments may be warranted to confirm this hypothesis, and in doing so it will be essential to carefully define the ER structure using a resident protein like calnexin. The fluorochrome DAPI may be useful for defining the boundaries of the nucleus while performing analysis of confocal microscopy images (Stuart and Cole, 2000). Additionally, quantitative analysis of multiple confocal images using densitometric analysis of a line segment drawn through the cell image would provide a better measure for determining the extent to which PM expression of hERG is reduced following STX1A coexpression (Um and Mcdonald, 2007). Finally, a higher resolution of hERG channel PM expression could potentially be obtained through the use of a hERG antibody directed towards an external epitope (for example, Alomone APC-109). Use of such an antibody would not require permeabilization, and would therefore reduce the background signal and the intense cytosolic fluorescence which we currently see in our images. Alternatively, use of an enhanced green fluorescent protein (EGFP) tagged hERG channel construct would significantly increase the resolution of confocal images because all hERG channels would be fluorescently tagged

(Claassen et al., 2008). The EGFP tag does not alter channel functionally when attached to the C- terminus, and this construct would allow for the imaging of live cells without the need for time- consuming preparation of fixed slides or cell-damaging permeabilization. We fully recognize the limitations of the qualitative observations obtained with this experiment, and so development of a more detailed result using the techniques describe may be warranted. However, these results are in agreement with an alternative assessment of hERG channel trafficking using Western blot followed by densitometric analysis.

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We further characterized the STX1A-mediated impairment of hERG channel trafficking by careful

Western blot analysis. The expression of 2 distinct protein bands corresponding to immature (core- glycosylated) and mature (complex-glycosylated) hERG allowed for the quantification of changes in total PM hERG protein expression. While the appearance of the 155 kDa hERG band signals successful protein maturation and trafficking out of the ER, it does not necessarily reflect trafficking to the plasma membrane (Gong et al., 2002). However, the ratio of 155 kDa/total hERG protein has been well established as an appropriate and reliable measure of hERG channel trafficking (Zhou et al., 1998b;

Phartiyal et al., 2008). Densitometric analysis revealed that a reduction in the 155 kDa/total hERG protein ratio was statistically significant and dose-dependent (see Fig. 21). This experiment also served to verify that while the amount of STX1A cDNA was sufficient to impair hERG channel trafficking, it did not completely abolish it. STX1A has also been found to reduce cell surface expression of KV1.1, KV2.1 and KV4.2 channels (Fili et al., 2001; Leung et al., 2003; Yamakawa et al., 2007).

Further experiments assessed the degree to which trafficking of the C-terminal truncated hERG constructs could be impaired by STX1A coexpression. STX1A significantly reduced hERG channel trafficking in HA-hERG-WT, ∆1120, ∆1045, and ∆1000 constructs, but not ∆960. We hypothesize that

STX1A functionally interacts with hERG channels upstream of residue 1000, producing its trafficking impairment effect. This finding is in agreement with our hypothesis that STX1A must interact with hERG close to the proximal cytosolic C-terminus in order to affect the voltage-sensitivity of hERG channel inactivation. hERG channels possess an ER-retention sequence at residues 1005-1007, characterized by an Arg-X-Arg motif, and subtle changes in channel structure near this motif have resulted in unmasking of the sequence and ER retention (Kupershmidt et al., 2002). It is possible that binding of STX1A upstream of this ER retention sequence results in its unmasking in WT and shorter truncation mutations, resulting in hERG trafficking inhibition. This may also explain why HA-hERG-

∆960 channel trafficking was not affected by STX1A, therefore warranting electrophysiological analysis of this truncation to assess whether STX1A modulates functionality of this construct.

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Trafficking impairment of hERG channels represents the most prevalent mechanism by which LQT2 mutations reduce native IKr (Anderson et al., 2006). Missense mutations in WT hERG can reduce or completely abolish channel maturation and trafficking by a variety of mechanisms. For example, missense mutations can impair normal hERG channel folding and assembly in the ER or may prevent export to the Golgi where complex glycosylation and sorting occurs prior to channel trafficking to the

PM (Furutani et al., 1999; Delisle et al., 2004). Some LQT2 mutants can form functional channels at the

PM when coexpressed with WT hERG, as mentioned previously. Trafficking-deficient LQT2 mutant channels generally produce reduced hERG current amplitude as well as alterations in channel gating kinetics or voltage-sensitivity if they successfully traffic to the PM (Anderson et al., 2006). LQT2 mutant hERG channels tend to be structurally diverse in the point mutations affecting normal hERG trafficking which are located throughout the channel sequence. In one study, 34 missense LQT2 hERG mutations were assessed, and 28 of these produced a trafficking-deficient phenotype as determined primarily by

Western blot analysis (Anderson et al., 2006). Although the locations of these missense mutations were dispersed throughout the hERG channel sequence, the majority of them were confined to highly ordered structures including α-helices and β-sheet domains (Anderson et al., 2006). Interestingly, 12 of these mutations were clustered between the S5 and S6 domains, most predominantly in the pore region of the channel. The characterization of these LQT2 mutations may shed some light on how

STX1A is affecting hERG in the present study, especially considering all of the similarities in the observed phenotype following coexpression. However, these types of comparisons would greatly benefit from the elucidation of the specific site of hERG-STX1A interaction.

Here, we have shown that STX1A-mediated impairment of hERG channel function is the result of a reduction in hERG PM expression as well as a reduction in channel open probability. STX1A- interaction is able to cause a functional modulation of the voltage sensitivity of hERG channel inactivation. As mentioned previously, residues important for inactivation of hERG channels are located in the pore region of the channel, and missense mutations of these residues are known to inhibit channel trafficking and appear to be highly clustered in this region. Therefore, the mechanism

-116- by which STX1A inhibits trafficking and enhances inactivation may be mediated by the same structurally-determined molecular interaction, potentially occurring adjacent to the S6 helices, thereby mechanically coupling to the pore region of the channel via this α-helix.

Several compounds have been recently shown to impair WT hERG channel trafficking, providing a new mechanism for drug-induced LQT syndrome and potentially providing insight into the mechanism by which impairment of channel trafficking may occur. For example, pentamidine is an antiprotozaol agent which does not cause hERG channel block at therapeutic concentrations.

However, a 2-day incubation of hERG-HEK 293 cells with the drug results in a marked reduction in whole-cell hERG currents and channel trafficking as measured by the reduction in the 155 kDa mature hERG Western blot band (Cordes et al., 2005; Eckhardt et al., 2005; Kuryshev et al., 2005). Arsenic trioxide has been demonstrated not only to directly block hERG channels, but to also strongly impair hERG channel trafficking, thereby reducing whole-cell hERG currents to an even greater extent (Drolet et al., 2004; Ficker et al., 2004). Celastrol, digitoxin (ouabain) and probucol also impair hERG channel trafficking, while only ceslastrol directly blocks hERG channel currents (Sun et al., 2006; Guo, J. et al.,

2007; Wang, L. et al., 2007). Of course these novel findings have important implications for drug screening of hERG channel trafficking impairment independently from that of hERG channel block

(Wible et al., 2005; Van Der Heyden et al., 2008). The mechanisms by which these compounds reduce hERG trafficking are poorly understood, and it is likely that different classes of compounds achieve their effects through independent mechanisms. For example, some compounds may disrupt interaction of immature hERG proteins with ER or Golgi chaperone proteins. Alternatively, some of these drugs may be capable of binding to the immature pore region of the channels preventing their proper folding or export through the ER-Golgi network.

hERG channel trafficking can be also impaired by inhibition of chaperone proteins and even by coexpression with the β-subunit MiRP1 (KCNE2). Normal hERG channel maturation involves careful regulation by quality control chaperone proteins including the heat shock proteins (see Chapter 1.3.5)

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(Walker et al., 2007). Inhibition of Hsp90 by the specific inhibitor geldanamycin causes inhibition of hERG channel maturation and ER retention (Ficker et al., 2003). The K+ channel β-subunit MiRP1 coimmunoprecipitates strongly with hERG, causing a reduced PM expression pattern (Um and

Mcdonald, 2007). MiRP1 impairs hERG channel trafficking by slowing down the rate of channel transport to the PM (Um and Mcdonald, 2007). Functionally, MiRP1 also inhibits hERG channel currents at the PM by accelerating the rate of channel activation and deactivation resulting in significantly reduced current amplitude (Abbott et al., 1999). It is possible that STX1A may inhibit hERG channel trafficking by disrupting normal chaperone protein interactions in the ER, or by slowing the rate of channel trafficking to the PM similar to MiRP1. More information regarding the molecular determinants of the structural-functional relationships underlying these complicated mechanisms is required.

The initial analysis of the dose dependence of STX1A coexpression produced one unexpected result – a very low transfection dose (10x less) of STX1A actually appeared to increase the density of the mature hERG protein band. This would suggest that STX1A may actually have a biphasic effect on hERG channel modulation. Remarkably, a similar observation was made by Fili and colleagues while investigating the regulation of KV1.1 by STX1A (Fili et al., 2001). At higher STX1A transfection levels, whole-cell current amplitude decreased, consistent with a decrease in channel trafficking to the PM.

At lower doses of STX1A transfection (approximately 8X lower) STX1A significantly increased KV1.1 currents with no detectable change in cell-surface expression. A more detailed electrophysiological analysis of the dose dependence of STX1A coexpression is required to further define this potential biphasic mechanism. Moreover, a lower level of STX1A expression would be consistent with a more physiologically relevant model, and as such, may be more reflective of the true nature of the endogenous interaction between the SNARE proteins and cardiac K+ channels. Additionally, this finding would likely satisfy the criticism that SNARE proteins are quite conformationally adaptable or

“sticky”, thereby increasing their propensity for non-specific interactions with other proteins (Jahn and

Scheller, 2006).

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Another interesting and unexpected finding was that STX1A actually increased the trafficking of HA- hERG-∆899, shifting the mature protein band to a higher molecular weight. This result is perplexing but it may provide insight into the potential biphasic mechanism by which STX1A could regulate hERG channels, increasing trafficking and restoring normal current amplitude under the right conditions. A shift in the protein band may indicate a very strong binding of a single hERG α-subunit and STX1A, such a shift may be possible considering the predicted molecular weight of STX1A (35 kDa). Of course further experiments are required to determine whether this is true. One simple experiment would be to perform Western blot analysis of the same sample and use an anti-STX1A antibody instead of the anti-HA antibody. Alternatively, one could repeat this experiment, and then strip the membrane and re-immunoblot it with anti-STX1A antibody. These preliminary results warrant electrophysiological analysis of the HA-hERG-∆899 truncation mutation without and with

STX1A coexpression. It would be interesting to see whether whole-cell HA-hERG-∆899 currents actually increase in response to STX1A coexpression with this mutant construct, and also to evaluate whether STX1A produces any changes in the voltage-sensitivity or gating kinetics of the channel, particularly the steady-state inactivation parameter.

5.4 STX1A-hERG binding experiments support functional data

In order to develop a more detailed understanding of the mechanism by which hERG and STX1A interact we investigated whether these proteins physically bind to one another. We hypothesized that the proteins directly interact, binding near the proximal cytosolic C-terminus located closest to the S6 domain based on the functional regulation of hERG channel trafficking and modulation of the voltage sensitivity of channel inactivation. We utilized GST pulldown and coimmunoprecipitation assays in order to develop the third aim of my project – to determine the site of hERG-STX1A binding.

GST pulldown of full length hERG and STX1A demonstrated that the two proteins can directly interact in vitro. We further tested two GST-fusion constructs comprising the HABC (amino acids 1-160) and the

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H3 (amino acids 191-256) domains of STX1A. hERG specifically bound to the STX1A-H3 domain which is known to serve as the active domain of the protein – the SNARE motif – and is involved in the formation of SNARE complexes during membrane fusion events (Jahn and Scheller, 2006). The HABC domain inhibits the function of the H3 SNARE motif by reversibly binding and forming a “closed” conformation (Misura et al., 2000). Previous work in our laboratory has demonstrated that the H3 domain interacts with KATP channels (Cui, N. et al., 2004), KV1.2 (Neshatian et al., 2007), KV2.1 (Lam et al.,

2005) and KV4.2 (Yamakawa et al., 2007), and so it was no surprise to see that this domain also strongly and preferentially interacts with hERG channels. Our lab has also tested the effects of a constitutively active “open-form” STX1A mutant (L165A/E166A) in which the HABC domain of STX1A is unable to flip over to block the C-terminal H3 domain (Dulubova et al., 1999). Open-form STX1A is a more potent inhibitor of KATP and KV2.1 channel function (Cui, N. et al., 2004; Leung et al., 2005). STX1A is an integral membrane protein and is characterized by a SNARE motif which lies on the intracellular side of the PM and is connected to the TM domain by a short rigid linker (Kiessling and Tamm, 2003). Therefore, it is possible the STX1A-H3 domain is perpendicularly oriented to the PM, exposing a considerable surface area of its α-helical structure to be available for interaction, and in the case of hERG with one of its large cytosolic N- or C-terminal domains.

Coimmunoprecipitation demonstrated that hERG and STX1A strongly bind to one another in our mammalian cell system. The use of hERG truncation mutations allowed us to further probe this interaction, and revealed that STX1A strongly interacts with hERG channels possessing large N- and C- terminal truncations. The hERG N-terminus, which contains the PAS domain, is approximately 395 amino acids in length. STX1A bound strongly with the largest N-terminal truncation hERG-∆2-354, which indicates that it is probably unlikely that the STX1A-H3 domain interacts at the N-terminus.

Alternatively, the hERG C-terminus possess the cNBD and starts at approximately residue 670, terminating at residue 1159. The largest C-terminal truncation mutation was HA-hERG-∆814 which also bound with STX1A. We hypothesize that the cytosolic STX1A-H3 domain interacts with the proximal cytosolic C-terminus between the residues 670 and 814. Such an interaction would support

-120- our functional results. In particular, an interaction between these residues could potentially alter the channel’s voltage-sensitivity to inactivation, causing the hyperpolarizing shift in steady-state inactivation. It could also account for a reduction in hERG channel trafficking, either by disrupting specific residues important for chaperone interactions located in the pore region, S6 domain or proximal C-terminus, or by possibly unmasking the downstream ER retention sequence and preventing proper folding of the channel.

Our laboratory has discovered that the site and mechanism of STX1A interaction with K+ channels varies greatly from channel to channel. This is in stark contrast to the highly conserved and well described interaction of SNARE proteins with CaV channels (Catterall, 1999; Atlas, 2001). The synaptic protein interaction site (synprint) motif located on the cytoplasmic II-III loop provides a common site for interaction of SNARE proteins including STX1A with CaV channels (Atlas, 2001; Zamponi, 2003). In contrast, the site of SNARE protein binding in K+ channels varies not only structurally but functionally from channel to channel. For example, we and other groups have previously reported that STX1A binds to the N-terminus of KV1.1 (Fili et al., 2001; Michaelevski et al., 2002) and KV4.2 (Yamakawa et al.,

2007). Interestingly, these channels are regulated by β-subunits which have also been demonstrated to interact with STX1A. In contrast, STX1A binds strongly to the C-terminus of KV2.1 (Leung et al.,

2003). Taken as a whole, while K+ channels lack a conserved synprint motif for SNARE protein interaction, this permits the exquisite and fine-tuned control of each channel type through the specific modulation of channel trafficking and function, governed by the unique structural characteristics of each protein.

There are limitations, however, to coimmunoprecipitation studies of this nature. In recently years, several groups have been highly critical of this approach to studying SNARE proteins because of their extreme conformational adaptability (Jahn and Scheller, 2006). Free SNARE motifs, such as the STX1A-

H3 domain, are known to be notoriously sticky, thereby producing a large number of non-specific bindings with other proteins. This means that SNARE proteins are susceptible to false positives with

-121- pulldown and immunoprecipitation assays. Over 100 binding partners have been reported for synaptic SNARE proteins (Jahn and Scheller, 2006). Binding of these proteins regulate the sorting and recycling of SNARE proteins during transport (Pennuto et al., 2003), control the recruitment of SNAREs into trafficking vesicles (Peden et al., 2001; Siniossoglou and Pelham, 2001), assist in the formation of docking complexes (Collins et al., 2005) and regulate the activity of the SNARE motif (Hu et al., 2002).

Determination of the in vivo relevance for SNARE protein interaction is critical for distinguishing between “test-tube phenomena” and genuine physiologically significant interactions.

5.5 Disruption of STX1A-mediated impairment of hERG channel trafficking

Rescue of numerous LQT2 trafficking-deficient mutations by reduced temperature and use of high- affinity channel blockers has been well established as a mechanism by which impaired hERG channel function can be restored in vitro (reviewed in Chapter 1.3.6) (Anderson et al., 2006; Robertson and

January, 2006). While the clinical applicability of these therapies is limited, they do provide a great deal of information regarding the mechanisms by which trafficking-impaired hERG channels operate.

For example, incubation of trafficking-deficient LQT2 mutants at reduced temperature boosts the expression of these difficult to express proteins by stabilizing the intermediate steps in the protein- folding pathway, thereby promoting their export from the ER (Delisle et al., 2004). With the knowledge that reduced temperature increases trafficking of membrane proteins like hERG, it has been experimentally determined that WT hERG channels yield the greatest expression of surface- associated protein when cultured at 30 °C (Chen et al., 2007).

In order to develop the fourth aim of my project, I attempted to disrupt the STX1A-mediated inhibition of hERG channel trafficking by growing cells at reduced temperature. This significantly restored hERG channel trafficking to the PM. In fact, both groups were statistically indistinguishable following Western blot and densitometric analysis of the 155 kDa/total hERG protein ratio. Based on this result, we hypothesize that the STX1A-interaction causing reduced hERG channel trafficking has

-122- been disrupted due to stabilization of the intermediate steps of protein-folding which may be inhibited by the STX1A interaction. We cannot, however, conclude that STX1A interaction has been abolished. In fact, this is unlikely given the strong binding affinities for these proteins. Further experimentation will be required to determine whether STX1A is still associated with hERG channels cultured at reduced temperature (coimmunoprecipitation) as well as electrophysiological characterization of these channels to determine whether whole-cell current amplitude has been significantly restored, and whether STX1A is still able to produce a hyperpolarizing shift in the midpoint of steady-state inactivation at this temperature.

The effects of the high-affinity hERG channel blocker E-4031 on STX1A trafficking-deficient hERG cells were also assessed using both electrophysiological and molecular biology methods. Peak hERG current amplitude and tail currents were significantly increased following 24 h E-4031 incubation and these currents were indistinguishable from hERG + E-4031 controls. Additionally, the hyperpolarizing shift in steady-state inactivation caused by STX1A was not affected, thereby providing evidence that while E-4031 may at least partially rescue channel trafficking, impairment of channel function at the

PM is unaffected. This implies that hERG and STX1A proteins do not lose their physical interaction during E-4031 rescue of hERG channel trafficking. To support this finding, Western blot analysis of HA- hERG cells cotransfected with STX1A illustrates that while densitometric analysis of the 155 kDa mature hERG band is not statistically different from control, there is certainly a trend toward increasing hERG channel maturation following E-4031 incubation. E-4031 binds to the inner vestibule region of hERG channels and acts to stabilize intermediate configurations of protein folding (Ficker et al., 2002). Binding and unbinding of E-4031 can only occur when a channel is in the open state, so the drug becomes easily trapped within the channel when the activation gate closes (Kamiya et al., 2006).

This may help explain why this drug is so potent, and how it may actually stabilize the immature protein from the inside out. While we cannot determine from these results whether E-4031 in fact disrupts the direct binding of STX1A to hERG channels within the ER-Golgi network, we can be sure that they are physically coupled at the PM based on electrophysiological analysis. It is more likely, as

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4031 acts to stabilize protein folding during channel maturation.

Interestingly, the results of Western blot analysis of hERG channel trafficking rescue do not seem to precisely mirror the electrophysiology data which demonstrates a statistically significant effect produced by E-4031 incubation. This may be primarily because of the lack of sensitivity of the Western blot technique. Alternatively, further experiments exploring this rescue effect could employ cell- surface biotinylation to specifically measure only hERG protein present at the PM. This technique was utilized by our lab previously while investigating the STX1A-mediated reduction in KV4.2 expression at the PM (Yamakawa et al., 2007). Cell surface biotinylation also allows for the direct densitometric analysis of PM hERG protein (not a ratio of 155 kDa/total protein) and therefore is not affected by fluctuations in cytosolic hERG expression which may occur as a result of ER-“trapping” or accumulation of immature or misfolded hERG channels. Alternatively, flow cytometry could be used to measure the fluorescent signal produced by PM hERG proteins labeled with an anti-hERG antibody directed towards an external epitope and conjugated to a fluorescent protein such as GFP (Takemasa et al.,

2008). Preparation of such a sample for flow cytometry would be relatively easy and would not require cellular permeabilization prior to incubation with antibodies. This technique would accurately generate a quantitative measure of the averaged relative fluorescence emissions of thousands of cells.

We hypothesize that these techniques would strongly correlate with our functional electrophysiology data and prove that E-4031 significantly increases the PM expression of hERG following STX1A- mediated inhibition of trafficking.

To further develop the project aim of disrupting STX1A-mediated hERG channel inhibition, a competitive binding assay could be developed similar to recent work published by Tsuk and colleagues (Tsuk et al., 2008). In this study, STX1A-mediated inhibition of KV2.1 channel function was abolished following expression of the full SNARE complex, comprised of STX1A, SNAP-25 and VAMP2.

Independently, these SNARE proteins uniquely regulate KV2.1 channel function; however, when the

-124- entire complex is present the individual SNAREs are recruited into full SNARE complexes which do not interact with the channel (Tsuk et al., 2008). This experiment could provide valuable insight into the relative strengths of channel versus SNARE complex binding and interaction and may represent a more physiologically relevant condition, in which the regulation of individual SNARE proteins results in excess of a particular component, thereby specifically altering the functionality of the channel in question.

Use of neurotoxins could also help to further probe the hERG-STX1A interaction. The clostridium neurotoxin, botulinum neurotoxin C1 (BoNT/C1) can proteolytically cleave STX1A inhibiting its function and subsequently promoting its degradation (Hayashi, T. et al., 1994; Dolly and Aoki, 2006).

Our laboratory has previously used BoNT/C1 to rescue the STX1A-mediated modulation of KV4.2 channel function (Yamakawa et al., 2007). In the context of this project, use of BoNT/C1 could cleave

STX1A proteins interacting with hERG channels at the PM and reverse the STX1A-moduation of hERG channel function. This would allow for an accurate calculation of the effect of reduced hERG channel trafficking on whole-cell current amplitude versus the effect of enhanced channel inactivation.

5.6 Endogenous expression and physiological relevance

The ultimate goal of this research program is to determine and establish the physiological role of

SNARE protein regulation of cardiac K+ ion channels. To this end, our laboratory has clearly demonstrated that a variety of channels including hERG are modulated in vitro by SNARE proteins including STX1A. However, the biological role for SNARE proteins in nonsecretory cardiomyocytes remains unclear. For this reason, the fifth and final aim of my research project is to begin to assess the endogenous expression of hERG channels and SNARE proteins in native cell systems. While this specific aim remains the least developed in my research program, it carries the greatest consequences for pursuing future investigation into this novel mechanism.

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Previously, our lab has identified the expression of STX1A in rat ventricular myocytes using confocal microscopy and Western blot analysis (Kang et al., 2004). More recently, the expression of two isoforms of STX1 in rat ventricular myocyte membrane using Western blot and mouse ventricular myocytes using confocal microscopy has been identified (Ng et al., 2008). Another group has also identified the expression of numerous SNARE proteins including STX1A in mouse atrial myocytes

(Peters et al., 2006). Here, I have demonstrated the expression of STX1A in mouse heart membrane isolated from whole hearts. This is not surprising as we have previously observed a high level of PM expression of STX1A in mouse ventricular myocytes (Ng et al., 2008). Additionally, our laboratory has detected other SNARE proteins in mouse heart and HL-1 cells including SNAP-25 and VAMP2 which are required for SNARE complex formation, as well as SNAP-23 and STX4 (unpublished data). Peters and colleagues suggest that SNARE proteins are responsible for the regulated exocytosis of atrial natriuretic peptide (ANP) in the endocrine heart (Peters et al., 2006). Furthermore, they demonstrate an isoform shift from STX4 and SNAP-23 to STX1A and SNAP-25 in the adult heart. Since we have demonstrated on numerous occasions the modulation of cardiac ion channel function including KATP

(Kang et al., 2004; Ng et al., 2008), KV4.2 (Yamakawa et al., 2007), KV4.3 (Ahmed et al., 2007) and now hERG channels by STX1A and SNAP-25 (see Appendix 2 for hERG + SNAP-25), we hypothesize that the regulation of K+ ion channel function by SNARE proteins becomes more physiologically significant in adults. Moreover, SNARE proteins may also be involved in the fine-tuning of ion channel expression and function related to excitation-contraction coupling, rhythmic control and regulation of ANP secretion (Ng et al., 2008).

Future experiments exploring the physiological interaction of hERG and STX1A in the heart are complicated by the need to prepare fresh cardiomyocytes on a daily basis for experimental assays including patch-clamp electrophysiology. Here, we have demonstrated that the HL-1 cell line derived from mouse atria (Claycomb et al., 1998), also expresses a variety of SNARE proteins including STX1A and ether-à-go-go related gene (ERG) proteins. Western blot analysis of ERG proteins in HL-1 cells revealed the expression of 3 bands. This is in agreement with work done by the Shrier laboratory with

-126- both experiments demonstrating protein bands at roughly the predicted 135 and 155 kDa molecular weights corresponding to immature and mature protein, respectively (Akhavan et al., 2003).

Additionally, a third band is visualized at approximately 80 kDa and this may correspond with the N- terminal truncation isoform ERG1b (Jones et al., 2004). Thus, HL-1 cells may possess ERG currents that more closely resemble native IKr in the mouse heart. These results are extremely exciting as they suggest that HL-1 cells may represent a physiological relevant model for studying the endogenous expression and interaction of SNARE proteins with cardiac K+ ion channels in a model system that is much easier to grow, maintain and use for experiments including patch-clamp analysis.

In order to probe the physiological role of SNARE proteins in the heart, it will be necessary to utilize native cardiomyocytes or HL-1 cells and to specifically disrupt the function of the SNARE proteins in these systems. As mentioned above, BotNT/C1 could be used to cleave STX1A, and following this several experimental methods could test the functional consequences of inhibition. For example,

Western blot analysis or cell surface biotinylation could assess changes in K+ channel PM expression, and electrophysiological recordings could assess changes in channel gating kinetics. In lieu neurotoxin use, small interfering RNAs (siRNA) could be designed and transfected into HL-1 cells to abolish function of SNARE proteins including STX1A.

These proposed experiments may have clinical implications as approximately 30-35% of patients diagnosed with LQTS cannot be linked to any of the known genes involved in this disorder (Crotti et al., 2008). It is possible that alterations in the expression patterns of SNARE proteins, especially a disproportionately large expression of STX1A relative to SNAP-25 and VAMP2 may inhibit hERG channel function leading to LQTS. This potentially novel mechanism underlying LQTS now seems increasingly feasible considering the growing list of mutations affecting channel-associated proteins that are not ion channels themselves.

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We believe that SNARE protein-mediated regulation of cardiac ion channels represents a novel biological mechanism that may have universally intrinsic implications for normal and diseased heart function. Downregulation of cardiac K+ channels including hERG has been implicated in cardiac disease in both human and animal models (Beuckelmann et al., 1993; Gidh-Jain et al., 1996). Careful investigation into the role of SNARE protein expression patterns and function in the diseased heart may prove to reveal mechanisms underlying conditions ranging from cardiac hypertrophy and hypertension to diabetes.

5.7 Conclusions

The main purpose of this research project was to explore the relationship between the voltage-gated cardiac ion channel hERG and the SNARE protein syntaxin 1A. Based on previous work carried out by our group examining the interaction of K+ channels with SNARE proteins, we hypothesized that STX1A is an important intrinsic regulator of hERG channel trafficking and gating. In order to assess this potential interaction, we devised a research program organized around 5 main aims.

Aim 1 – To perform electrophysiological assessment of the hERG-STX1A interaction. STX1A drastically reduces whole-cell hERG current amplitude following coexpression in mammalian cells. This is at least partially achieved because of STX1A-mediated modulation of the voltage-sensitivity of hERG channel inactivation. STX1A induces a significant hyperpolarizing shift in the mid-point voltage of steady-state inactivation, but does not, however, affect the kinetics of channel activation, deactivation, fast inactivation or recovery from inactivation.

Aim 2 – To evaluate STX1A-induced changes in hERG trafficking, surface expression and localization.

Immunocytochemistry and confocal microscopy revealed that coexpression of hERG and STX1A results in a decreased PM expression of hERG protein. These two proteins colocalize with periplasmic distribution in highly-dense intracellular organelles. Western blot analysis reveals that STX1A

-128- significantly reduces hERG channel maturation as measured by the intensity of the 155 kDa mature hERG protein band. The STX1A-mediated reduction in hERG channel maturation is dose-dependent and may be biphasic in nature with STX1A actually promoting hERG channel trafficking at low concentrations. Additionally, use of C-terminal truncation mutations revealed that STX1A significantly reduces trafficking of HA-hERG-∆1120, ∆1045 and ∆1000, but not ∆960. Functional interaction of hERG and STX1A occurs upstream of residue 1000. Interestingly, maturation of HA-hERG-∆899 actually increased following STX1A coexpression, but this mechanism requires further study. These experiments demonstrate that the large reduction in hERG current amplitude following STX1A coexpression is primarily due to a reduction in the number of functional channels expressed at the PM.

Additionally, the primary mechanism by which STX1A inhibits hERG channel function is by inhibiting channel maturation and trafficking.

Aim 3 – To determine the site of hERG-STX1A binding. GST pulldown assays demonstrated that hERG and

STX1A can bind in an overexpression system. Moreover, hERG protein preferentially interacted with the STX1A-H3 domain which contains the active SNARE motif. Coimmunoprecipitation demonstrated that hERG and STX1A bind in our mammalian cell system. Use of truncation mutations illustrated that

STX1A was capable of binding to hERG channels with large N- and C-terminal truncations. Based on these results and the structure of the STX1A H3 domain and the large hERG-channel C-terminus we concluded that STX1A interacts with hERG at the proximal cytosolic C-terminus. Interaction at this site has functional consequences for hERG voltage-sensitivity to inactivation and trafficking.

Aim 4 – To disrupt STX1A-mediated changes in hERG channel function or expression. Incubation of hERG cells coexpressed with STX1A at reduced temperature for 24 h resulted in a significant increase in mature hERG protein, thereby disrupting the trafficking-inhibition effect of STX1A. Alternatively, incubation of these cells with the hERG channel blocker E-4031 resulted in significantly increased hERG current amplitude as determined by electrophysiological analysis. Western blot analysis of hERG channel maturation showed a trend towards trafficking rescue, however, this was not statistically

-129- significant, possibly due to the limited resolution of this method. Rescue of channel trafficking by reduced temperature and high-affinity drug block is believed to occur by stabilization of immature proteins and lowering of the critical free energy barrier during folding and assembly, thereby promoting N-linked glycosylation and processing through ER quality control checkpoints (Akhavan et al., 2003; Robertson and January, 2006). Interestingly, the STX1A-mediated hyperpolarizing shift in steady-state inactivation was still present following rescue of channel trafficking, indicating that this inhibition of channel function is a secondary mechanism of hERG channel inhibition.

Aim 5 – To evaluate the endogenous expression of hERG and STX1A in cardiac myocytes and HL-1 cells. Use of membrane protein obtained from mouse cardiomyocytes allowed us to demonstrate the expression of ERG protein and several SNARE proteins including STX1A. Additionally, we demonstrated the endogenous expression of these proteins in HL-1 cells which may serve as an uncomplicated yet physiologically relevant system for further study of the SNARE protein regulation of cardiac ion channels.

To summarize, we have demonstrated that STX1A inhibits hERG channel function by two mechanisms

(Fig. 32). Firstly, it impairs channel maturation and trafficking to the PM and this is the primary mechanism by which whole-cell hERG currents are impaired. Secondly, STX1A physically interacts with hERG channels at the PM and alters the voltage-sensitivity of inactivation, further inhibiting channel function. The STX1A-mediated inhibition of hERG channel function represents a novel mechanism for regulation of the predominant repolarizing current of the mammalian heart. STX1A may be an essential intrinsic regulator of cardiac K+ channel function and this has important clinical implications for heart disease.

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Figure 32. STX1A impairs hERG channel function - mechanism

Macroscopic IhERG is the product of the total number of channels at the PM (N), the probability that a channel will open (PO), and the single-channel conductance (i). Coexpression of STX1A and hERG proteins result in a remarkable inhibition of hERG channel function. The primary mechanism by which impairment of hERG channel function occurs is by the STX1A-mediated inhibition of hERG channel maturation and trafficking to the PM. (This likely involves the disruption of normal protein folding and assembling in the ER and possibly complex glycosylation at the Golgi. This leads to the ER retention of immature hERG with STX1A proteins.) The secondary mechanism by which hERG channel function is inhibited is via a direct interaction of these proteins at the PM, resulting in the production of a hyperpolarizing shift in the voltage-sensitivity of steady-state inactivation of hERG channels. Together, these mechanisms result in a significant reduction in macroscopic hERG current amplitude and reduced channel function.

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5.8 Summary of recommendations for future experiments

Throughout Chapter 5, I have noted limitations and made recommendations for future experiments based the results obtained and information from complementary investigations. Here, I have summarized two new aims for this research program indicating essential experiments to be performed for their development.

New Aim #1 – To better define the mechanism by which STX1A regulates hERG channel function

1. Perform immunocytochemistry and confocal microscopy by utilizing antibodies directed towards

a hERG external epitope, use of ER marker proteins or EGFP-hERG constructs to improve our

understanding of PM expression and localization of hERG channels.

2. In order to enhance the limited resolution of WB analysis of hERG channel trafficking following

temperature or E-4031 dependent rescue, perform cell surface biotinylation or flow cytometry to

quantify alterations in PM expression of hERG channels. Additionally, use of additional structurally

diverse compounds such as thapsigargin or cisapride for assessment of hERG channel rescue may

provide insight into the structural determinants of this mechanism.

3. To determine the extent to which hERG channel inhibition is the result of trafficking impairment or

functional regulation use neurotoxins to specifically cleave STX1A and perform patch-clamp

electrophysiology to measure whole-cell hERG currents. This will allow for the calculation of the

proportional effect of each inhibitory mechanism thereby negating the need for single-channel

recordings.

4. In order to assess the effect of STX1A on the rate of hERG channel trafficking, degradation and

channel turnover, perform pulse-chase analysis of hERG protein maturation.

5. We made the interesting observation that low doses of STX1A may increase hERG channel

maturation. This warrants further molecular biology and electrophysiological assessment of low

dose STX1A transfection in order to determine whether STX1A actually has a biphasic affect on

hERG channel function.

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6. Finally, we made the interesting observation that STX1A increases the trafficking of the HA-hERG-

∆899 mutation. Further investigation of this finding may provide evidence for the mechanism of

STX1A-impairment of hERG channels. Western blot analysis should be used to test the mature

hERG protein band which has shifted to a higher molecular weight in order to determine if STX1A

has is in fact tightly bound to hERG. Additionally, this finding may warrant electrophysiological

recordings.

New Aim #2 – To explore the physiological importance of SNARE proteins in the heart

1. In order to firmly establish the endogenous role for regulation of hERG channels by STX1A,

perform coimmunoprecipitation of these proteins in native cardiomyocytes and HL-1 cells.

2. Determine the physiological importance of SNARE proteins including STX1A in cardiomyocytes

and HL-1 cells by abolishing their function through the use of siRNA or neurotoxins. Then assess

ion channel trafficking and maturation using Western blot analysis, cell-surface biotinylation,

confocal immunofluorescence, flow cytometry and patch-clamp electrophysiology.

3. Establish the importance of SNARE complex formation by coexpressing in vitro STX1A, SNAP-25

and VAMP2 and determining whether their interaction inhibits the individual regulation of ion

channels including hERG by STX1A. Then, use siRNA or neurotoxins to abolish the function of one

of these SNARE complex forming proteins in primary or HL-1 cells. Assess whether the absence of

or relative abundance of SNARE proteins affects ion channel trafficking or functionality using

Western blot analysis, confocal immunofluorescence, flow cytometry and patch-clamp

electrophysiology.

4. Test the expression patterns of the SNARE proteins in cardiac disease models including cardiac

hypertrophy, hypertension and diabetes. Obtain cardiomyocytes from animal models of these

diseases and determine whether alterations in SNARE protein expression can be correlated with

alterations in ion channel function in these diseases.

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Appendix 1

Preliminary results: Interaction of hERG and STX1A coexpression in tsA-201 cells

Voltage-gated K+ channel (hERG) recordings of single tsA 201 cells were performed using the whole- cell configuration of the patch clamp technique. Recordings were made using a HEKA EPC-10 amplifier and Pulse software. Pipettes were pulled from 1.5 mm borosilicate glass capillary tubes

(World Precision Instruments, Sarasota, FL) using a programmable micropipette puller (Sutter

Instrument, Novato, CA). Pipettes were heat polished and resistances were obtained ranging from 2-4

MΩ when filled with a solution containing (in mM): 140 KCl, 1 MgCl2, 5 EGTA, 10 HEPES, 5 ATP (pH 7.2 with KOH). The bath solution contained (in mM): 140 NaCl, 4 KCl, 1 CaCl2, 1 MgCl2, 10 glucose, 5 HEPES

(pH 7.4 adjusted with NaOH). Once whole-cell configuration was established, the cell was held at -80 mV and subjected to various experimental protocols. All recordings were performed at room temperature (~22°C).

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Appendix 1a: Effect of STX1A on hERG current amplitude

A) Whole-cell hERG currents were elicited by 3 s depolarizing pulses from -80 to +60 mV in 10 mV increments from a holding potential of -80 mV. Tail currents were elicited by a step repolarizing pulse to -60 mV for 3 s. Robust hERG current expression is observed, displaying its unique rapid entry in C-type inactivation at more depolarized potentials and large outward tail currents. B) Coexpression of STX1A resulted in reduced hERG current amplitude and the more noticeable absence of C-type inactivation. C) Current-voltage relationship measured at the end of the 3 s depolarizing pulse from cells expressing hERG alone (n=18) and hERG + STX1A (n=16). A significant reduction in outward current is observed at potentials positive to -30 mV. D) Voltage dependence of hERG activation measured from the tail currents. STX1A significantly reduced peak conductance but had no influence on the midpoint of activation (-9.1 ± 1.1 mV vs. -9.4 ± 1.9 mV).

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Appendix 1b: Effect of STX1A on hERG rate of activation

A) Activation time constants were measured by fitting the currents elicited by 3 s depolarizing pulses using the monoexponential equation. Summarized data for cells expressing hERG alone (n=16) and hERG + STX1A (n=10). STX1A caused a significant acceleration in the rate of activation between -10 mV and +60 mV (p < 0.05).

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Control

Appendix 1c: Effect of STX1A on hERG deactivation

A) Whole-cell tail currents were elicited from -140 mV to +20 mV for 5 s following a 1 s depolarizing pulse to +60 mV in cells expressing A) hERG alone or B) hERG + STX1A. Deactivation rates were measured by fitting tail currents to a biexponential function as shown. C) Deactivation fast and slow time constants (τ) for hERG (n=16) and hERG + STX1A (n=9) shown in panel A and B. STX1A has no significant effect on the rate of hERG deactivation.

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Appendix 1d: Effect of STX1A on hERG steady-state inactivation

A) Whole-cell hERG channel steady-state inactivation was measured using a triple-pulse protocol. Cells were depolarized for 3 s to +60 mV (holding potential of -80 mV), followed by conditioning pulses from -140 mV to +20 mV for 20 ms, and then a test pulse to +20 mV for 500 ms. Only the first 80 ms is shown for the test pulse. B) Summarized steady-state inactivation data for hERG and hERG + STX1A. STX1A produced a significant hyperpolarizing shift in the midpoint steady-state inactivation curve from -29.9 ± 4.1 mV (n=12) to -51.5 ± 4.8 mV (n=6) with no change in the slope factor (20.6 ± 1.0 mV vs. 19.2 ± 1.2 mV).

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Appendix 2

Preliminary results: Interaction of hERG and SNAP-25 coexpression in tsA-201 cells

Appendix 2a: SNAP 25 inhibits whole-cell hERG currents

SNAP-25 was coexpressed in TSA 201 cells in a 2:1 ratio (hERG: SNAP-25 or STX1A) in order to electrophysiologically assess a potential interaction between the two SNARE proteins and hERG. (A) Peak hERG currents were significantly reduced from 136.1 ± 7.6 pA/pF (hERG, n=18, +20 mV) to 64.3 ± 12.0 pA/pF (hERG + SNAP-25, n=9, +10 mV) (p<0.01). (Cotransfection with STX1A: 42.0 ± 6.1 pA/pF (hERG + STX1A, n=15 at +20 mV)). B) Peak tail currents at +60 mV were also reduced from 99.1 ± 8.8 pA/pF (hERG, n=18) to 65.3 ± 10.5 pA/pF (hERG + SNAP25, n=9) (p<0.05). (Cotransfection with STX1A: 43.6 ± 8.6 pA/pF (hERG + STX1A n=15)). Although there was no change in slope factor, there was a significant hyperpolarizing shift in the midpoint of steady-state activation from -9.1 ± 1.1 mV (hERG n=18) to -16.3 ± 1.6 mV (hERG + SNAP25 n=9) (p<0.01). (STX1A : -9.39 ± 1.9 mV). C) Steady-state inactivation analysis reveals that there is no change in slope or midpoint of steady-state inactivation following cotransfection with SNAP25, despite a prominent hyperpolarizing shift in the midpoint of steady-state inactivation in the presence of STX1A (hERG: -29.9 ± 4.1 mV, n=12; hERG + STX1A: -51.5 ± 4.8 mV, n=6; hERG + SNAP25: -26.1 ± 7.2 mV, n=5).

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Appendix 2b: SNAP-25 reduces mature hERG channel trafficking

The inhibition of hERG channel maturation by SNAP25 was assessed by Western blot analysis. Coexpression with varying concentrations of SNAP25 cDNA resulted in a decreased mature complex glycosylated hERG band at 155 kDa following transfection with 0.5 μg SNAP25 (2 hERG: 1 SNAP25 ratio). Lower transfection ratios did not yield significant reductions in mature hERG channel expression. α/β tubulin was used as a loading control (lower panel). B) Quantification of channel trafficking was assessed by measuring the fraction of mature/total hERG protein by densitometric analysis of the immunoblots (n=3). SNAP25 significantly reduces 155 kDa hERG/ total hERG from 0.25 ± 0.03 to 0.10 ± 0.03 when 0.5 μg of SNAP25 cDNA is transfected.